PROTEIN ANALYSIS IN VITRO

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purification protein electrophoresis immunoassays detecting the interaction sites of DNA-binding proteins measuring protein-protein interactions protein conjugation incorporation of synthetic amino acids into proteins nonribosomal peptide synthesis (NRPS) analyzing mAb specificity protein engineering increasing serum half-lives of proteins

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Protein analysis

• Web resources
• Protein engineering glossary
• Centre for Protein Engineering (CPE) at Cambridge University
• Peptide libraries to determine the substrate specificity of protein kinases
• proten purification
• Preparation of active proteins from inclusion bodies
• dot blot technique : technique for detecting, analyzing, and identifying proteins, similar to the Western blot technique but differing in that protein samples are not separated electrophoretically but are spotted through circular templates directly onto the membrane or paper substrate.
• protein electrophoresis : the separation of ionic solutes based on differences in their rates of migration in an applied electric field.
• microelectrophoresis : electrophoresis in which migrating particles are observed by light microscopy
• isoelectric focusing (IEF) / electrofocusing : electrophoresis in which the protein mixture is subjected to an electric field in a gel medium in which a pH gradient has been established; each protein then migrates until it reaches the site (or focus) at which the pH is equal to its isoelectric point (the pH at which it has no net charge)
• moving boundary electrophoresis : the original method of electrophoresis, in which the movement of the solvent is unrestricted and all of the particles of a species move at the same rate, maintaining a sharp boundary which can be optically monitored.
• zone electrophoresis : any of several methods of electrophoresis in which an inert support medium holds the molecules as they migrate in the conducting medium, thus preventing convection and diffusion.
• paper electrophoresis : an older method of zone electrophoresis in which the support medium is paper, mainly used to separate serum proteins.
• cellulose acetate electrophoresis : a method of zone electrophoresis in which the support medium is a sheet of cellulose acetate; used mainly in clinical chemistry to analyze or purify serum proteins.
• capillary electrophoresis (CE)
• high performance capillary electrophoresis (HPCE)
• gel electrophoresis : a type of zone electrophoresis in which the support medium is a gel, in the form of tubes or a thin slab; it is usually composed of agarose, polyacrylamide, or starch and is so named. Specific types are used to separate certain classes of molecules on the basis of change, size, or both.
• starch gel electrophoresis : an older form of gel electrophoresis using a hydrolyzed starch support matrix to separate macromolecules, particularly proteins.
• agarose gel electrophoresis : a type of gel electrophoresis with agarose as the support medium; used extensively to separate proteins, lipoproteins, nucleic acids, and other substances.
• polyacrylamide gel electrophoresis (PAGE) : gel electrophoresis using a polymerized polyacrylamide matrix to separate molecules on the basis of size, charge, or both; usually used to separate proteins or sequence nucleic acids. Gels are ... :
• usually discontinuous
• disc electrophoresis : a method of polyacrylamide gel electrophoresis involving discontinuous (hence the name) gel layers. A discontinuity in pore size and pH between the layers is used to prevent diffusion and maximize separation of the components.
• but may be a single layer
..., and are either ...
• nondenaturing to examine native molecules
• denaturing
• SDS–polyacrylamide gel electrophoresis (SDS-PAGE) : a type of polyacrylamide gel electrophoresis in which the anionic detergent sodium dodecyl sulfate (SDS) is used to denature the sample proteins into linear monomers, rendering their charge proportional to their length so that migration is a function of size.

Web resources :
• SDS-polyacrylamide gels: standard Laemmli protocol
• Ethanol-based SDS-PAGE Coomassie Blue staining
• Bench protocol for silver staining SDS-polyacrylamide gels
Web resources :
• Tricine-polyacrylamide gels: for high resolution of small proteins
• two-dimensional gel electrophoresis (2D-GE) : a method for improved separation of complex mixtures of molecules by subjecting the support medium to electrophoresis in 2 directions, usually at right angles to each other; e.g., isoelectric focusing (IEF) followed by SDS-PAGE => proteins are separated by the independent parameters of isoelectric point in the first dimension and by molecular weight in the second
• 2D difference gel electrophoresis (2D-DIGE) : a fluorescence-based method that increases the power of the proteomics techniques by allowing 2 different protein samples tagged with 2 distinct fluorescent dyes to be run on the same gel
After IEF in the presence of urea and a nonionic detergent, the IEF gel is equilibrated in sodium dodecyl sulfate (SDS) to prepare the proteins for SDS-PAGE. The method described here^ref uses carrier ampholytes to form a pH gradient in a long, thin (1.2-mm) focusing gel composed of a low percentage (2.7%) of acrylamide and containing 9.5 M urea and 2% Nonidet P-40 to maintain protein solubility. After IEF, the gel is briefly equilibrated in SDS and placed directly onto the top edge of a second-dimension slab gel. Because the time between the end of IEF and the start of SDS-PAGE is only a few minutes (thereby minimizing diffusion), the spots are highly resolved and nearly round in shape^{ref1, ref2}.
• first dimension: IEF
• gather the IEF gel tubes and set up the IEF casting apparatus. A handy casting device^ref fills up to 15 gel tubes automatically. The IEF tubes are placed upright in the tube containing the IEF solution, and this assembly is immersed at a controlled rate into a cylinder of water, displacing the solution upward into the tubes and maintaining constant polymerization temperature.
• thaw an aliquot of IEF gel solution and add 10% ammonium persulfate to initiate polymerization.
• cast the IEF gels so that they are filled to within 1-2 cm from the top and allow them to polymerize at 30 °C for 60 min.
• assemble the IEF tubes in the first-dimension IEF apparatus and fill the appropriate chambers with upper (0.1 N NaOH) and lower (0.01 M H₃PO₄) electrode buffers.
• add 3 ml of overlay buffer to each tube, using a microliter syringe or a thin pipette tip. Connect the electrode wires and prefocus the gels to establish the pH gradient. Start the current at 100 A per gel and continue until the voltage has reached 1,000 V.
• using the needle of a microliter syringe or an elongated pipette tip, apply the sample (5-15 ml) directly to the gel surface, displacing the overlay buffer upward.
• carry out IEF overnight for 19,000 volt-hours, that is, 18 h at 1,000 V and 1 h at 2,000 V (for 20-cm-long tubes), or according to manufacturer's instructions for different formats. Increasing the voltage to 2,000 V for the last 1 h will help sharpen the bands.
• remove the IEF gels from the first-dimension apparatus and lay them sample end forward on a bed of ice. The urea in the gels should not precipitate; if it does, this indicates impurities in the urea or other solutions.
• second dimension: equilibrium and electrophoresis
• cast the slab gels as for one-dimensional SDS-PAGE (Laemmli or tricine procedure) and assemble the gel cassettes into the second-dimension electrophoresis apparatus. Fill the electrode chambers with the appropriate electrode buffer.
• push the IEF gels from the glass tubes, one at a time, into a small tray of equilibration buffer. As the tube emerges, dip the acidic end briefly into 1% bromophenol blue to mark it. Shake at 37 °C for 1 min.
• immediately transfer the IEF gel into the upper electrode buffer chamber. Under the buffer, align the IEF gel on the notched glass plate with the marked acidic end on the left. Roll the IEF gel into the space between the plates using a small spatula and forceps and seat the gel snugly onto the slab gel without stretching.
• attach electrode wire, put safety shields in place and apply the current. The total time between the end of IEF and the beginning of SDS electrophoresis should ideally be just a few minutes.
• carry out slab gel electrophoresis as for one-dimensional gels.
• immunoelectrophoresis (IEP) : a technique combining protein electrophoresis and double immunodiffusion; proteins are separated by agarose gel electrophoresis; then specific (polyclonal) antisera are placed in a trough cut parallel to the protein track, and the proteins and antibodies are allowed to diffuse through the gel, the proteins diffusing radially from their electrophoretic placement and the antibodies diffusing perpendicularly from the trough, resulting in a distinct elliptical precipitin arc for each protein detectable by the antisera
• counter electrophoresis / counterimmunoelectrophoresis (CIE) / countercurrent immunoelectrophoresis : one-dimensional double electroimmunodiffusion; a technique in which antibody and antigen are placed in separate wells in an agar plate and driven toward each other by an applied electric field, because the gel is buffered at a pH between the isoelectric points of the antigen and antibody. It is more sensitive and faster than double immunodiffusion and is particularly useful for antigens that diffuse slowly in the gel
• crossed immunoelectrophoresis (Laurell's first technique) : a combination of protein electrophoresis and rocket immunoelectrophoresis; protein antigens are separated by agarose gel electrophoresis; then a strip containing the separated antigens is cut out and placed in a trough in a gel containing antiserum, and an electric field perpendicular to the trough is applied, producing a “rocket” precipitin pattern for each antigen
• rocket immunoelectrophoresis (Laurell's second technique) : one-dimensional single electroimmunodiffusion; a technique in which antigen is placed in a row of wells in an agar plate containing antiserum and an electric field perpendicular to the line of wells is applied; this drives the antigen through the gel, forming a spike or “rocket” precipitin pattern trailing away from each well. The length of the rocket is proportional to the amount of antigen placed in the well
• immunofixation electrophoresis (IFE) is a 2-stage process consisting of agarose gel electrophoresis of the patient serum  followed by fixation of proteins and application of monospecific antisera to IgG, IgA, IgM, k and l chains (one per track), which undergo a single immunodiffusion reaction. It has replaced IEP as the method of choice for identification and characterisation of monoclonal immunoglobulins in serum, urine and cerebrospinal fluid. IFE allows ease of interpretation, fast results (< 2 hours) and has excellent sensitivity to low levels of monoclonal immunoglobulin (Ig) as may be found in cases of early multiple myeloma / MGUS. Samples are normally screened using serum protein electrophoresis, those with abnormal results are further investigated using Freelite and/or IFE.
• Tiselius apparatus : an apparatus for the electrophoretic separation of the proteins of blood serum, plasma, and other body fluids.
• electrophoretogram / electropherogram / electrophoregram : the record produced on or in a supporting medium by bands of material which have been separated by the process of electrophoresis
• immunoblot affinity purification : the immunoblotting method has evolved from early stages when antibodies were used to 'stain' polyacrylamide gels directly^{ref1, ref2} to more versatile methods using replica techniques, in which the separated polypeptides are transferred to nitrocellulose membranes, chemically activated paper or nylon sheets. Although there are several variations on this basic theme, the most common and effective is electrophoretic transfer to nitrocellulose sheets^{ref1, ref2}. The separated proteins can then be probed with antibodies; this variation of the technique is known as immunoblotting (or western blotting). The membrane can also be probed with specific ligands, such as DNA, protein, small molecules (for example, heparin or GTP) or even whole cells. Electrophoretic transfer can be achieved either in a tank^ref or in a semidry apparatus^ref, in which the buffer volume is reduced to filter paper pads. The following protocol presents the original method used for tank blotting^ref
• transfer of proteins to a solid support :
• 1| Cut filter paper (Whatman 3MM or Bio-Rad Blot absorbent paper) and the membrane to the dimensions of the gel (or portion of it) to be transferred.
• 2| Soak both membrane (if using polyvinylidene fluoride (PVDF), wet with methanol first and wash with deionized water) and filter papers in appropriate transfer buffer for 10 min after slowly immersing them at a 45° angle. Towbin transfer buffer: 25 mm Tris-Cl, 192 mm glycine (pH 8.3), 20% methanol; do not adjust the pH. The transfer buffer can be chilled to 4 °C before transfer if desired.
• 3| Rinse the gel in water (or transfer buffer) briefly. Leaving the gel in this solution too long will remove SDS from the proteins, decreasing their solubility.
• 4| Set up the transfer apparatus. For tank transfers it is best to assemble the gel-membrane sandwich in transfer buffer.
• Ensure that the fiber pads for the tank transfer are well soaked in buffer and that air bubbles are removed.
• Assemble the materials in order from cathode side to anode side: fiber pad, two sheets of filter paper, gel, membrane, two sheets of filter paper, fiber pad.
• Clamp this sandwich together in the transfer cassette according to the apparatus being used. Half fill the tank with buffer and insert the cassette; then quickly fill the tank with buffer to above the cassette height.
• Transfer at 100 V (250 mA) for 60 min; circulate the buffer in the tank with a revolving stir bar.
These conditions serve as a guide, for example, for a Bio-Rad Mini Gel Transfer; for other systems, see manufacturer's recommendations.
• 5| Turn off the electric current and disconnect the leads. Disassemble the transfer apparatus from the top downward, removing each layer in turn.
• 6| Transfer the gel to a tray containing Coomassie brilliant blue, and stain it to determine whether the electrophoretic transfer was complete.
• 7| Optional: Stain the membrane filter with Ponceau S or india ink. India ink should be used only if the blot is to be probed with a radiolabeled antibody or probe.
• detection by blocking and probing with antibody
• 8| After electrophoretic transfer, rinse the blotted membrane with deionized water, then block nonspecific binding sites with blocking buffer (5% bovine serum albumin (BSA) or 5% nonfat dried milk in Tween-20 Tris-buffered saline (TTBS; for nonfat dried milk, warming the solution will assist dissolution). TTBS: 0.05% Tween-20 in TBS; TBS: 20 mm Tris-Cl, 500 mm NaCl (pH 7.5)) at 15-25 °C for 1 h with agitation. Remove the buffer
• 9| Rinse the blot twice with TTBS for 1 min each time.
• 10| Incubate the blot in the appropriately diluted primary antibody solution in incubation buffer (1% BSA in TTBS) at 15-25 °C for 1 h with agitation. Remove the solution.
• 11| Rinse the blot with TTBS at 15-25 °C. Rinse once rapidly; then wash once for 15 min and twice for 5 min.
• 12| Incubate with secondary antibody (alkaline phosphatase or horseradish peroxidase (HRP) linked so that it can be used for colorimetric or chemiluminescent detection) in TTBS at the recommended dilution (usually 1:5,000) at 15-25 °C for 30 min with agitation. Remove the solution.
• 13| Rinse as in Step 11, then rinse twice more in TBS for 5 min each time.
• 14| Perform the appropriate protocol for detection, for example, for radiolabeled, colorimetric or chemiluminescent detection. For colorimetric detection, you may use the Bio-Rad protocol using alkaline phosphatase with BCIP/NBT (5-bromo-4-chloro-3-indolyl phosphate/nitroblue tetrazolium); for chemiluminescent detection, the ECL protocol using horseradish peroxidase (HRP)-conjugated secondary antibody.
• Western blot technique : a technique for analyzing and identifying protein antigens: the proteins are separated by electrophoresis in polyacrylamide gel, then transferred (“blotted”) onto a nitrocellulose membrane or treated paper, where they bind in the same pattern as they formed in the gel. The antigen is overlaid first with antibody, then with anti-immunoglobulin or protein A labeled with a radioisotope, fluorescent dye, or enzyme :
• transfer proteins from gel to membrane (nitrocellulose of polyvinylidene fluoride (PVDF)): gloves should be worn when handling gels or blot membranes.
• pour enough transfer buffer (or CAPS buffer) into a container so that half of the assembled cassette will be 0.5 cm below the surface of the buffer.
• after the gel has finished running, carefully remove the front glass plate and remove the stacking gel. Remove a small portion of the upper left-hand corner of the running gel to mark its orientation during the transfer process.
• wet the membrane with transfer buffer and carefully lay it across the exposed running gel, making sure that air bubbles between the gel and membrane are excluded. In the same fashion, add two layers of pre-wetted filter paper to the membrane. Pre-wet one of the cassette sponges and add to the stack. Top with the outer half of the cassette itself and carefully turn the entire stack of glass plate, gel, membrane, filter paper, sponge and cassette over. Place the stack in the container with transfer buffer.
• carefully remove the second glass plate from the gel. Using the same process, make sure that no air bubbles have formed between the gel and the membrane.
• keeping the stack generously wet, top the gel with 2 layers of pre-wet filter paper, wet sponge and the remaining half of the cassette unit. Snap the cassette unit together firmly.
• place the cassette into the transfer unit so that the current will flow through the gel onto the membrane. This would be with the gel closer to the black (-) cathode and the membrane closer to the red (+) anode.
• add transfer (or CAPS) buffer to the transfer unit according to manufacturer's directions. Transfer over night with a setting of 20V / 40 mA or for 3 hours at 70V/160 mA. The transfer process needs to take place under cold temperatures to prevent the gel from sticking to the membrane. This can be accomplished either by using a water cooling core in the transfer unit or placing the entire unit at 4°C. Please follow the manufacturer's recommendations.
• at the end of the transfer period, disconnect the power connections and disassemble the cassette. All of the Coomassie Blue-dyed bands should now be on the membrane leaving the gel fairly clear.
• staining the blot (optional) :
• briefly with water, rinse any bits of gel off the blot then incubate it in Ponceau S solution for 1 minute. Rinse in water until the background is reduced.
• place the membrane on a transparency sheet and photocopy the blot for a permanent transfer record.
• rinse the blot several times in TBS-Tween to remove the stain.
• blocking the blot :
• gently shake the blot for 2 hours in 5% non-fat dry milk/TBS/Tween at room temperature or at 4°C on a rocker.
• either pour off blocking buffer or transfer the blot to a sealing bag for probing.
• probing the blot:
• dilute the primary antibody in 5% non-fat dry milk/TBS. Add to the blot, seal the container and gently rock for 2 hours at room temperature or over night at 4° C. If using a bag for the incubation, eliminate all air bubbles before sealing.
• remove the primary antibody and wash the blot 3 x 5 minutes in TBS/Tween. In some instances, the primary antibody can be saved for additional blot procedures.
• dilute the secondary antibody in 5% non-fat dry milk/TBS. Add to the blot, seal and incubate as before.
• remove the secondary antibody and wash the blot 3 x 10 minutes in TBS/Tween.
• detection / chemiluminescence:
• cut 2 transparency sheets to fit into the film cassette.
• mix oxidizing and luminol reagent (from Dupont NEN) in a 1:1 ratio.
• pour onto a piece of parafilm and lay the blot on the pool of solution for 1 minute.
• insert the blot into the cassette and top with the remaining sheet of transparency, making sure that all bubbles are eliminated.
• insert film and close cassette. Start with a 1 minute exposure and adjust from there. The signal will decay over time. If no bands are found, the blot can be rinsed and stored in PBS/Tween overnight for repeated blocking and probing.
• stripping and reprobing blots : works well with chemiluminescence and radioactive detection, but is not useful in colorimetric detection due to the production of an insoluble reaction product formed as a results of that detection method.
• briefly rinse the blot in water.
• wash the blot 2 x 10 minutes in TBS/Tween.
• re-block in 5% milk/TBS/Tween for one hour at room temperature.
• add stripping buffer to the blot, seal the container andi ncubate for 30 minutes at 50° C. Occasionally agitate the container for best results.
• wash the blot 2 x 30 minutes followed by 2 x 10 minutes in TBS/Tween.
• if desired (optional), can incubate the blot in the detection reagent and expose to film to ensure that no residual detection antibody remains. In order to check for the presence of primary antibody, the blot would have to be incubated with secondary antibody and detection reagent to verify complete removal.
• block the blot for 2 hours in 5% milk/TBS/Tween at room temperature before probing with the desired primary antibody again.
A Western blot may provide some advantage in situations where your antibody may cross react with other proteins.  For example, since you can identify your protein of interest by molecular weight, you can determine the activity of the antibody against the protein vs background noise (non-specific binding). The main advantage of the ELISA is that the protein is still intact ie in its quaterniary / tertiary structure and so can be analysed from both structural and functional integrity (eg with cell based assays), whilst the western will denature you protein and only provide you with limited information regarding its characteristics.
• Southwestern blot technique : a technique analogous to Southern blot technique but in which proteins are separated electrophoretically, transferred to a nitrocellulose filter, and probed with radioactive or otherwise labeled fragment of DNA; performed to detect expression of a specific DNA binding protein, such as a transcription factor.
• radioligand assay : any assay procedure that uses radioisotopic labeling and biologically specific binding of reagents, such as
• radioimmunoassay (RIA)
• radioreceptor assay : a radioligand assay in which a radiolabeled hormone is used to measure the concentration of specific cellular receptors for the hormone in tissue specimens, an example being radioassay of estrogen receptors in breast tissue
The interactions of antibodies with ...
• insoluble antigens => agglutination reactions are very sensitive, they often exhibit inhibition at high Ab concentrations and are carried out at physiological pH and ionic strength (this is clearly a requirement for RBCs but is also a requirement for bacterial agglutination which is inhibited at low ionic strength and occurs in an antibody-independent manner at high ionic strength). Agglutination plays a very important role in the diagnosis of bacterial infections and in blood typing. The latter has been conducted for many decades with natural (IgM) antibodies. IgM are strong haemoagglutinins. In contrast other antibodies not only fail to produce haemoagglutination but positively inhibits it (blocking antibodies). Anti-Rh antibodies are a classic example : these antibodies either bind in a monovalent maner or they bind in a divalent manner to the same cell. They can be revealed by using an antiglobulin antiserum (Coombs reagent). The simplicity and sensitivity of the haemoaglutination reactions can be considerably widened in the so-called passive haemo-agglutination. RBCs, latex, etc. readily absorb many polysaccharides and protein Ags can be coupled covalently using bifunctional reagents.
• antiglobulin reaction : the agglutination of particles (usually erythrocytes) that have been (1) sensitized by the adsorption of soluble antigen, (2) treated with antibody to that antigen, and (3) treated with antiserum to the serum globulin of the animal species that produced the antibody
• Coombs' antiglobulin test (AGT) : a test for the presence of nonagglutinating antibodies against red cells that uses antihuman globulin antibody to agglutinate red cells coated with the nonagglutinating antibody
• the direct antiglobulin test detects antibodies bound to circulating red cells in vivo. It is used in the evaluation of autoimmune and drug-induced hemolytic anemia and hemolytic disease of the newborn
• the indirect antiglobulin test detects serum antibodies that bind to red cells in an in vitro incubation step. It is used in typing of erythrocyte antigens and in compatibility testing (cross-match)
• antiglobulin consumption test : a test for serum antibodies against cellular antigens. Cells are incubated with the serum sample and then with antiglobulin; any serum antibody that binds to the cells will take up antiglobulin. The amount of antiglobulin consumed is determined by testing the supernatant with antibody-coated red cells; the amount of agglutination is inversely proportional to the antiglobulin consumption.
• conglutination reaction : a characteristic agglutination reaction obtained by a mixture of conglutinin, cells (e.g., bacteria or red cells), fresh complement, and a cell-specific immune serum from which the agglutinins have been removed by absorption. See conglutinin.
• conglutinating complement absorption test (CCAT) : a test resembling the complement fixation test, using as the indicator of antigen-antibody reaction the disappearance of conglutinin activity.
• mixed agglutination reaction : agglutination of a mixture of different cell types by antibody directed against an antigenic determinant present on all of the cells.
• agglutination test (for presence of antibody) : cells containing antigens to a given antibody are mixed into the solution being tested; agglutination indicates presence of the antibody
• latex agglutination or fixation test (for presence of antibody): a type of agglutination test in which antigen to a given antibody is adsorbed to latex particles and mixed with a solution to observe for agglutination of the latex
• soluble antigens : the complexes formed between a mAb and its Ag or the complexes formed between a polyclonal antibody and a hapten are soluble. However the complexes between polyclonal antibodies or mixtures of monoclonal antibodies and Ags often lead to the formation of an insoluble precipitate, the so-called precipitin reaction / precipitation (it occurs only with multivalent antigens and is dependent on electrolyte concentration, pH, temperature, and the relative concentrations of antigen and antibody, the amount of precipitate formed increasing to a maximum and then decreasing as the relative antigen concentration is increased). Such reactions can occur both in liquid media and in semisolid (gel) media and both media are widely exploited for applications in laboratory research and practice. The amount of precipitate that is recovered when an increasing amount of Ag is added to a constant amount of Ab in solution follows a typical pattern. The amount of Ab precipitated increases with the amount of Ag (zone 1 or prezone or Ab excess zone) up to maximum (zone 2 or equivalence zone), after which further increase in the amount of Ag leads to a decrease in the amount of Ab precipitated (zone 3 or postzone or Ag excess zone). Ab-Ag complexes in solution can form aggregates of various size and that when the volume of such aggregates exceeded a critical value, precipitation occurred [the sedimentation coefficient (S) of a particle is directly proportional to its volume (V), the difference between its density (r) and that of the solvent (r₀) and the gravitational field (g) : S = V (r-r₀) g]. Such latex theory explains well the different Ab/Ag ratios and the fact that mAbs cannot precipitate Ag. After a precipitate has formed, its complexes can dissociate and re-equilibrate with new Ag. If the latter is present in sufficient excess, soluble complexes will prevail and the initial precipitate will gradually dissolve. Thus, in principle, precipitation reactions are reversible. In practice, in the majority of the cases they are are not and a striking illustration of this point is the so-called Danysz phenomenon: when an equivalent amount of diphteria toxin is added to an anti-toxin serum, the mixture is non-toxic; however, if the same amount of toxins is added to the same amount of antiserum in aliquots and at intervals (of 30 mins, for example) the mixture is toxic. This is readily explained by the fact that the first addition of the toxin leads to the formation of complexes with a high Ab/Ag ratio and very slow dissociation rates (days or months). As a result, insufficient Ab is left to react with new toxin added in relatively short periods of time.
• precipitin test : any serologic test based on a precipitin reaction
• there are anomalous precipitation reactions (flocculation) in which precipitation is observed in a narrow range of Ab/Ag ratios. Such reactions probably require the presence of high-affinity, non precipitating antibodies (monogamous binders) which must be saturated with Ag before remaining, 'conventional' Abs can bind and precipitate the additional Ag.
• bentonite flocculation test : any agglutination test using antigen adsorbed on particles of bentonite; when the antigen is added to serum containing specific antibodies, flocculation occurs
• Ramon flocculation test : to a series of tubes containing a constant amount of toxin, e.g., diphtheria toxin, antitoxin is added in increasing amounts; when a zone of flocculation appears, the tube showing it contains a completely neutralized mixture of toxin and antitoxin. The first tube in which flocculation occurs is taken as the end-point.
• rapid plasma reagin (RPR) test : a flocculation test for syphilis using unheated serum and a modified VDRL antigen containing choline chloride and charcoal particles, enabling macroscopic identification of the flocculation; widely used for screening.
• automated reagin test (ART) : a modification of the rapid plasma reagin (RPR) test for use with automated analyzers; used in clinical chemistry
• microprecipitation test : a precipitin test in which a minute quantity of the serum is employed
• radioimmunoprecipitation assay (RIPA) : immunoprecipitation conducted with radiolabeled antibody or antigen.
• immunodiffusion / gel diffusion : any technique involving diffusion of antigen or antibody through a semisolid medium, usually agar or agarose gel, resulting in a precipitin reaction. Precipitin lines or bands form where the concentrations of antigen and antibody are serologically equivalent.
• diffusion : the spontaneous movement of molecules or other particles in solution, owing to their random thermal motion, to reach a uniform concentration throughout the solvent, a process requiring no addition of energy to the system
• single diffusion : immunodiffusion in which either the antibody or antigen remains fixed and the other reactant diffuses through it.
• single radial diffusion / radial immunodiffusion (RID) in 1 dimension (Oudin) : a quantitative immunodiffusion technique in which the antigen solutions are placed in wells cut in an agar plate containing antiserum; the area or diameter of the precipitin ring around an unknown solution is compared with the rings of a serial dilution of a standard antigen solution to determine the amount of antigen present in the unknown. in which an Ag solution is left to diffuse over an antiserum that is 'trapped' in a gel and a precipitate forms in the agar in regions corresponding to the equivalence zone
• double diffusion : immunodiffusion in which both the antigen and antibody diffuse through the medium toward each other.
• double diffusion in one dimension (Oakley-Fulthorpe technique) : antiserum is placed in a test tube and overlaid with agar, which is allowed to solidify, and antigen is layered over the agar. Precipitin lines form where the concentrations of each antigen and antibody are equivalent
• double diffusion in 2 dimensions (Ouchterlony technique) : double diffusion in which antigen and antiserum are placed in wells cut in an agar plate; antigen solutions to be compared are placed in wells equidistant from the antiserum well. A large number of geometric arrangements are possible that enable analysis of Ag mixtures (reactions of identity, partial identity, non identity). Modifications of these techniques allow quantification of Ag or Ab. For example, if a uniform concentration of Ab is incorporated in the gel and Ag is applied in a well, the distance at which the precipitin ring forms, depends on the concentration of Ag in the well (Mancini). Thus, with appropriate standards, the concentration of Ag in unknown samples can be calculated from a calibration curve of [Ag] vs d (diameter) or a (area). Further modifications combine gel diffusion and precipitation with electrophoretic separation of the Ag. This can be carried out in order to analyse complex Ag mixtures (Grabar) or in order to decrease the time required in order to achieve precipitation and increase sensitivty of the Mancini-type procedure (Laurell). 3 principal types of reaction may occur, each identified by a characteristic pattern of precipitin lines, indicating the extent to which the antigen samples a share antigenic determinants :
• reaction of partial identity : a reaction pattern seen in double diffusion in two dimensions: one of the precipitin lines between the antigen wells and the antiserum well stops at the point of intersection, whereas the other continues past it, indicating that the antigen samples have some, but not all, antigenic determinants in common
• reaction of identity : a reaction pattern seen in double diffusion in 2 dimensions: the precipitin lines between the antigen wells and the antiserum well stop at their point of intersection, indicating that the antigen samples are identical
• reaction of nonidentity : a reaction pattern seen in double diffusion in two dimensions: the precipitin lines between the antigen wells and the antiserum well cross, indicating that the antigen samples have no antigenic determinants in common
Web resources :
• Analysis of protein by immunoprecipitation
• Immunoprecipitation with soluble antibodies
• fibrinogen assay : an assay for the level of fibrinogen in a plasma sample; a thrombin reagent is added to the diluted sample and the time needed to form clots is compared with that for a reference containing a known amount of fibrinogen
• 4-point assay : an assay based on a mixture of 1 doses of test material and 2 doses of standard material.
• Immunoassay : any of several methods for the quantitative determination of chemical substances that utilize the highly specific binding between an antigen or hapten and homologous antibodies, including ...
• antigen-antibody reaction : the reversible binding of antigen to homologous antibody brought about by the formation of weak bonds between antigenic determinants on antigen molecules and antigen binding sites on immunoglobulin molecules. Certain effects, e.g., precipitation and agglutination reactions and viral neutralization, result from the binding itself; other effects, e.g., complement activation and opsonization, result from conformational changes that occur in immunoglobulin molecules with antigen binding and enable them to interact with complement proteins and Fc receptors on phagocytes.
• toxin-antitoxin reaction : the antigen-antibody reaction between a toxin and antitoxin.
• antigen capture assay : one used to identify minute quantities of antigen in solution: large quantities of antibody against the desired antigen are fixed to a solid support matrix, over which the solution is passed. The antigen is retained by the matrix and can be identified by reaction with labeled antibody.
• E rosette assay : an assay for human T lymphocytes based on the existence of a specific receptor on T cells for a sheep red blood cell membrane antigen. Peripheral blood lymphocytes are incubated with sheep red cells. T cells are surrounded by a ring of red cells—an E (erythrocyte) rosette—and are counted using a hemacytometer.
• EAC rosette assay : an assay for human B lymphocytes using complement receptors, a B cell marker. Peripheral blood cells are mixed with ox red blood cells, IgM antierythrocyte antibody, and complement deficient in C5 (to prevent red cell lysis). The antibody-and-complement-coated erythrocytes (EAC) form rosettes with B cells, which are counted using a hemocytometer
• D-dimer assay : an immunoassay for the fibrin degradation product D dimer. Levels are elevated in deep venous thrombosis, acute myocardial infarction, pulmonary embolism, unstable angina, and disseminated intravascular coagulation (DIC). Called also D-dimer test.
• enzyme immunoassay (EIA) : any of several immunoassay methods that use an enzyme covalently linked to an antigen or antibody as a label, the 2 most common being
• enzyme-linked immunosorbent assay (ELISA) : any EIA utilizing an enzyme-labeled immunoreactant (antigen or antibody) and an immunosorbent (antigen or antibody bound to a solid support). A variety of methods (e.g., competitive binding between the labeled reactant and unlabeled unknown, or a sandwich technique in which the unknown binds both the immunosorbent and labeled antibody) may be used to measure the unknown concentration.
• an enzyme conjugated to antibody generates a colored reaction product from a colorless substrate. It is a safe, inexpensive, and convenient means of detecting ...
• specific antibody (indirect ELISA)
• antigen is adsorbed to wells of a plastic plate in a monomolecular layer
• an irrelevant protein is used to block remaining free sites on the plastic
• gelatin
• PBS/5%NCS
• casein (2% w/v)
• BSA
• skim milk
• tryptone peptone
• egg yolk
• patient's serum is added following washing
• enzyme-conjugated anti-immunoglobulin is added followed by washing
E.g. :
• anti-dsDNA test : an EIA that uses native dsDNA as an antigen to detect and monitor increased serum levels of anti-DNA antibodies, a sign of systemic lupus erythematosus; used in both detection and management of disease.
• antigen (sandwich ELISA)
• double-antibody sandwich enzyme-linked immunosorbent assays (DAS-ELISA)
• double antibody sandwich indirect ELISA (DASI-ELISA)
• substrate is added and color reaction is quantified by absorbance using an automated plate reader
• fast activated cell-based ELISA (FACE) kits (source : Active Motif) provide you with a better alternative to Western blots and sandwich ELISAs for studying protein phosphorylation because the cell-based format of FACE allows you to grow, fix and assay your cells all in a single 96-well plate. Not only does this eliminate the need for cell extracts, gels and radioactivity, but it helps to keep protein modifications to a minimum, so you can assess the exact protein state in the cell at your chosen time point. And, the format makes it easy to screen large number of compounds (< 2 hrs of hand-on time). FACE kits provide 2 antibodies to compare both phosphorylated and native protein levels, as well as secondary antibody, developing reagent and a cell quantification solution for normalizing cell numbers.
Web resources :
• ELISA home page
• ELISA Protocol and Troubleshooting : tech handbook from Chemicon
• enzyme-multiplied immunoassay technique (EMIT) : trademark for a homogeneous (single phase) enzyme immunoassay which utilizes the change in enzyme activity of an enzyme-labeled hapten that occurs on binding with antibody to determine the amount of unlabeled hapten (the unknown) present in a biologic specimen
• sounding immunoassay
• dot immunobinding assays (DIBA) :
• fluorescent or fluorescence immunoassay (FIA) / fluoroimmunoassay / immunofluorescence assay (IFA) : any immunoassay using fluorochrome-labeled antibody or antigen; classified as
• heterogeneous fluorescence immunoassay : a fluorescence immunoassay that has more than one phase, requiring separation of the bound label (antigen-antibody complex) from the free labeled immunoreactant by physiochemical methods.
• homogeneous fluorescence immunoassay : a fluorescence immunoassay that is single-phase and uses a change in fluorescence (such as quenching or increased polarization) accompanying antigen-antibody binding to measure the amount of bound label without separation.
• immunofluorescence / fluorescent antibody technique : an immunofluorescence technique in which antigen in tissue sections is located by homologous antibody labeled with fluorochrome (the single-layer technique) or by treating the antigen with unlabeled antibody followed by a second layer of labeled antiglobulin which is reactive with the unlabeled antibody (double-layer technique). Variations include direct, indirect, inhibition, and complement staining techniques.
• direct immunofluorescence (DIF) / direct fluorescent antibody technique (DFAT)
• the primary antibody is tagged with the fluorochrome
• indirect immunofluorescence (IIF)
• binding of unlabelled primary antibody to the substrate is visualized with a fluorochrome-conjugated anti-immunoglobulin preparation. Advantages are: increased sensitivity since several fluorescent secondary antibodies bind to each primary antibody simplicity since each primary antibody need not be fluoresceinated. If for diagnostic aims, specificity is low due to cross-reactivity with antigens from non-pathogenetic species, but it can be increased by
• diluiting serum (1:200) : Ab titre during infection by a pathogenetic species is usually much higher than during colonization by a non pathogenetic species.
• add the non-pathogenetic species to buffer cross-reactive Abs and let only specific Abs bind the intended, pathogenetic one
E.g. :
• lupus band test : an immunofluorescence test to determine the presence and extent of immunoglobulin and complement deposits at the dermal-epidermal junction of skin specimens from patients with systemic lupus erythematosus.
Web resources :
• Conjugation of mAbs to fluorescent dyes
• Troubleshooting nonspecific immunofluoresent labeling at Albert Einstein College of Medicine, Yeshiva University
• radioimmunoassays (RIA) : a highly sensitive and specific assay method that uses the competition between radiolabeled and unlabeled substances in an antigen-antibody reaction to determine the concentration of the unlabeled substance; it can be used to determine antibody concentrations or to determine the concentration of any substance against which specific antibody can be produced
• competitive protein-binding assay : a RIA in which labeled and unlabeled ligands compete for sites on a carrier that has the same avidity for both; the concentration of unlabeled ligand is inversely proportional to the amount of labeled ligand bound.
• immunoradiometric assay (IRMA) : a variant of RIA in which the antigen being measured reacts directly with radiolabeled antibody
• direct and quantitative comparison of MHC-presented peptides in pairs of samples, such as transfected and untransfected, tumorous and normal or infected and uninfected tissues or cell lines : isotopically labeled reagents to quantify by mass spectrometry the ratio of peptides from each source. The first method involves acetylation and is both fast and simple. However, higher peptide recoveries and a finer sensitivity are achieved by the second method, which combines guanidination and nicotinylation^ref
• radioimmunosorbent test (RIST) : a highly sensitive radioimmunoassay for measuring the total IgE antibody concentration in serum; the serum sample is reacted with radiolabeled IgE and anti–human IgE antibody coupled to an insoluble support. The amount of labeled IgE remaining bound to the immunosorbent varies inversely with the amount of (unlabeled) IgE present in the sample
• radioallergosorbent test (RAST) : a test used to measure specific IgE antibodies in serum. Allergen extract is coupled to a solid matrix (paper, cellulose particles); this immunosorbent is reacted with serum and washed and then reacted with radiolabeled anti–human IgE antibody and washed. Uptake of the labeled antibody is proportional to the level of specific serum IgE antibodies to the allergen. RAST is used as an alternative to skin tests to determine sensitivity to suspected allergens.
Web resources : Intensifying screens
• antibody microarray^ref: when it comes to enzymes, protein abundance does not necessarily translate into protein activity: post-translational modifications such as phosphorylation can dramatically alter enzyme function. Researchers traditionally have overcome this discrepancy with activity-based protein profiling (ABPP), which screens enzyme activity using probes that bind covalently to active sites. Most ABPP experiments employ in-gel fluorescence scanning to detect probe-labeled enzymes, but this method has several problems, including slow throughput, limited sensitivity, and poor resolution. In Cravatt's method, enzymes are labeled with fluorescent-tagged probes and incubated with a glass slide containing spatially segregated spots of antibodies targeting a particular enzyme class. The slide is washed and the active, probe-labeled proteases that remain bound are quantified by fluorescence scanning of the microarray^ref. In experiments with proteases, the microarray proved roughly 30 to 60 times more sensitive than gel-based methods. In gels, the enzymes you study migrate together, and because they're often similar in mass, the likelihood they overlap in migration patterns is pretty high and not as resolvable. The microarray gives you really nice resolution because it's mass-independent.  Moreover, the microarray consumed 10 times less material than gels per experiment, which should be of great value for the analysis of clinical samples, which are often of limited quantity. The microarray has been sublicensed by Activx Biosciences for both drug discovery and diagnostics development. While the new microarray is an excellent proof of principle, microarray-compatible antibodies are available for only a tiny percentage of the proteome. It may take a while before you have an array sophisticated enough to catch up with its enormous potential as a powerful tool. Nevertheless, probes for more than a dozen enzyme families have been created, focusing on the most popular 30 to 40 proteases or kinases used for diagnostic human-tissue profiling. The protease probes could be mixed with extracts and hybridized to microarrays composed of small molecules from a combinatorial library. Mass spectrometry or other methods could identify the protease that binds to any particular spot. That spot would then, in essence, function in the same way as an immobilized antibody. Those kinds of experiments could be done by some time in 2005.

Web resources : companies offering array-based products and services : Advalytix, Affibody, Affymetrix, Agilent, Applied Biosystems, Applied Precision, ARRM, Beckman Coulter, Becton Dickinson, Biacore, BioGenex, Bio-Rad, BMG Lab Technologies, Cambridge Peptides, Cell Signaling Technology, Dualsystems Biotech, GE/Amersham Biosciences, Genetix, Genomic Solutions, Greiner Bio-One, Hypromatrix, Invitrogen/Protometrix, KBiosystems, Molecular Devices, MorphoSys/Antibodies by Design, Nalge Nunc International, NanoPlex Technologies, NextGen Sciences, New England Peptide, EMD Biosciences/Novagen, Panomics, Pepscan Systems, Peptide Specialty Laboratories, PerkinElmer Life Sciences, Pierce, Proteome Systems, Qiagen, RoboDesign, Scienion, Sigma-Aldrich, Stratagene, SuperArray Bioscience, TeleChem International, Whatman/Schleicher & Schuell Bioscience, Zeptosens, Zyomyx
Web resources : Introduction to antibodies : tech handbook from Chemicon
Companies offering protein purification products : Agarose Bead Technologies; Agilent; Amersham Biosciences (now part of GE Healthcare); Applied Biosystems; Beckman Coulter; BD Biosciences Clontech; Biochrom Labs Inc.; Bio-Rad; BioScience Beads; Brinkmann; Calbiochem; Caliper Life Sciences; Center for Integrated Biosystems; Chemicon; Ciphergen; Dynal Biotech; EMD Biosciences; Eppendorf; Fluidigm; GenScript; Gyros; Invitrogen; J.T. Baker; MDS Inc.; Millipore; Molecular Probes; NCI/SAIC Protein Purification Group; New England BioLabs; Novagen; Paragon Bioservices; PhyNexus; Pierce Biotechnology; Polysciences, Inc.; Protana; QBIOgene; QIAGEN; SEPIAtec; Sigma-Aldrich; Stratagene; SuNyx; Tecan; Tosoh Biosep; Trenzyme; Vivascience; Whatman
• Detecting the interaction sites of DNA-binding proteins
• electrophoretic mobility shift assay (EMSA) / gel retardation assay : one of the most sensitive methods for studying the DNA-binding properties of a protein, can be used to deduce the binding parameters and relative affinities of a protein for one or more DNA sites or for comparing the affinities of different proteins for the same sites^ref. It is also useful for studying higher-order complexes containing several proteins, observed as a 'supershift assay'. EMSA also can be used to study protein- or sequence-dependent DNA bending^ref. In an EMSA, or simple 'gel shift', a ³²P-labeled DNA fragment containing a specific DNA site is incubated with a candidate DNA-binding protein. The protein-DNA complexes are separated from free (unbound) DNA by electrophoresis through a nondenaturing polyacrylamide gel. The protein retards the mobility of the DNA fragments to which it binds; thus, the free DNA migrates faster through the gel than does the DNA-protein complex. An image of the gel reveals the positions of the free and bound ³²P-labeled DNA. Procedure^ref :
• preparation of the gel and binding reaction :
• 1| Prepare a 40-ml 4.5% native acrylamide gel (using 1 to 1.5-mm spacers):
• acrylamide mix (30%; 29:1 acrylamide:bisacrylamide) : 6 ml
• 5x Tris-borate/EDTA (TBE) buffer : 4 ml
• glycerol (20% vol/vol) : 2 ml
• water : 28 ml
• ammonium persulfate (100% solution in water) : 300 ml
• N,N,N',N'-tetramethylethylenediamine (TEMED) (add just before pouring the gel) : 30 ml
Prerun the 'gel for 2 h at 10 mA. Large protein complexes greater than 1 MDa in mass are difficult to resolve and migrate quite slowly in the native polyacrylamide gels. For analyzing larger complexes, use either agarose or acrylamide-agarose composite gels.
• 2| Set up binding reactions in 0.5-ml siliconized microcentrifuge tubes:
•  recombinant protein (0.5-100 ng) : 1.00 ml
• ³²P-labeled DNA template (ideally 1 fmol) : 1.00 ml
• poly(dI:dC) (1 mg/ml) : 0.20 ml
• bovine serum albumin (BSA : 50 mg/ml) : 0.25 ml
• dimethylsulfoxide (DTT) (0.1 M) : 0.10 ml
• MgCl₂ (0.1 M) : 0.75 ml
• buffer D (20 mM HEPES-KOH (pH 7.9), 20% glycerol (vol/vol), 0.2 mM EDTA, 0.1 M KCl, 0.5 mM phenylmethylsulfonyl fluoride (PMSF), 1 mM DTT Add PMSF and DTT just before use) : 6.70 ml
• water : to 10 ml
The amounts of recombinant protein and the DNA template must be titrated. MgCl₂ is typically omitted when crude extracts are used as the source of protein. For binding measurements on short DNA sites, perform these reactions with double-stranded oligonucleotides. The effective length limit for restriction fragments is generally around 250 base pairs (bp).
• 3| Incubate the reaction at 30°C (incubate at 15-25 °C or on ice for crude extracts) for 1 h.
• 4| Load the samples directly (with no dye) onto the gel. Carefully layer the mix onto the bottom of the well and observe the schlieren line form at the glycerol-buffer interface.
• Into one empty lane, load 5 ml of 10X DNA loading buffer with dyes for use as a marker to determine how far the products have traveled in the gel.
• the mobility shift assay
• 5| Run the gel for desired time at 10 mA; for a 30-bp fragment, allow the bromophenol blue dye to migrate about two-thirds of the way down the gel.
• 6| When the electrophoresis run is complete, carefully pour out the buffer into the sink and remove the gel from the apparatus. Remove the comb and split the plates, leaving the gel attached to one plate. To fix the gel continue with Step 7; otherwise proceed to Step 8.
• 7| (Optional) Fix the gel in gel fixing solution (200 ml methanol, 100 ml acetic acid, 700 ml water) at 15?25 °C for 15 min.
• 8| Place the gel on two sheets of Whatman 3MM paper. Cover the other side of the gel with Saran Wrap and dry on a gel dryer at 70 °C for 1 h.
• 9| Expose the gel to autoradiography film or phosphorimager screen overnight.
• chromatin immunoprecipitation (ChIP) : protein-DNA interactions are captured in vivo by cross-linking DNA-binding protein to other binding sites (BSs). The DNA is fragmented and co-immunoprecipitated with the protein of interest using a specific antibody. The isolated DNA fragments are amplified by non-specific PCR, labelled with a fluorescent dye (yellow circles, usually Cy3 or Cy5) and identified by hybridization to a DNA microarray that contains probes for all genes, as well as for intergenic regions.
• genome-wide mapping technique (GMAT) combines chromatin immunoprecipitation (ChIP) of hyper-acetylated histones, with serial analysis of gene expression (SAGE) technology and could provide an alternative to ChIP-on-a-Chip techniques^ref. ChIP-on-a-Chip is limited to what's on the microarray, mostly open-reading frames not bound by regulatory molecules : microarray sequences are often the whole gene in one spot, providing no information on where in the gene it is modified. Also, ChIP on a chip quantification depends on hybridization efficiency, which is not an issue in GMAT. The acetylation level of the yeast genome is 20 to 50 times higher than the human genome, which is 250 times bigger than the yeast. Theoretically, only 10 times more sequencing would cover it
• acetylation microarrays combine ChIP and DNA microarrays to determine rapidly the level of histone acetylation throughout the entire genome. By identifying the regions at which acetylation is increased in the absence of, for example, a particular HDAC, acetylation microarrays allow for mapping the sites of action of the enzyme on a genome-wide scale. To do this, yeast cells are crosslinked with formaldehyde in vivo and subsequently disrupted mechanically. The chromaint released from cells is sheared by sonication to an average size of 400-500 bp for optimal resolution of the mapping experiments. Chromatin fragments from cells mutated for a specific HDAC (HDACD) and their isogenic wild type counterparts are immunoprecipitated using highly specific antibodies raised against individual sites of histone acetylation. DNA from enriched chromatin fragments is purified after reversing the histone-DNA crosslinks. The purified DNA is amplified linearly by PCR and labelled with a fluorophore (Cy3 or Cy5). Probes from both sets of labelled DNA are then combined and hybridized to a DNA microarray containing either intergenic regions, ORFs or both. For a given genomic region of the microarray, the ratio of the normalized fluorescent intensity between the 2 probes indicates whether the acetylation level of the analysed lysine residue is affected in the mutant strain
Web resources : Chromatin Immunoprecipitation (ChIP) at STKE [free registration required !]
• bacterial one-hybrid system^ref
• protein binding microarrays (PBMs) allows rapid, high-throughput characterization of the in vitro DNA binding–site sequence specificities of transcription factors in a single day^ref.
Web resources : Mapping Protein/DNA Interactions by Cross-Linking, Paris: Institut National de la Santé et de la Recherche Médicale; 2001 [free at NCBI BookShelf !]
• Measuring protein-protein interactions
• pairwise protein interactions
• coimmunoprecipitation : many protein-protein associations that exist within the cell remain intact when a cell is lysed under nondenaturing conditions. Thus, if protein X is immunoprecipitated, then protein Y, which stably associated with X, may also precipitate. Coimmunoprecipitation is most commonly used to test whether two proteins of interest are associated in vivo, but it can also be used to identify interacting partners of a target protein. In both cases, the cells, labeled with [³⁵S]methionine, are collected and lysed under conditions that preserve protein-protein interactions. The target protein is specifically immunoprecipitated from the cell extracts, and the immunoprecipitates are fractionated by SDS-PAGE. Coimmunoprecipitated proteins are detected by autoradiography and/or by western blotting with an antibody directed against that protein. The identity of interacting proteins may be established or confirmed by Edman degradation of tryptic peptides. Some early examples of this method include the use of antibodies to viral antigens to determine the host cellular proteins that interact with these viral transforming oncoproteins. 2 interacting proteins of particular note are the tumor suppressor proteins p53 and pRB^ref. This protocol was used to identify pVHL-associated proteins; conditions should be optimized for the protein of interest^{ref1, ref2, ref3, ref4}.

Procedure
• Preparation and precipitation of the lysate
• 1| Wash 30 10-cm plates of the appropriate cells (a total of 6 x 10⁷ cells) in phosphate-buffered saline. Scrape each plate of cells into 1 ml of ice-cold EBC lysis buffer (50 mM Tris (pH 8), 120 mM NaCl, 0.5% NP-40, 50 mg/ml leupeptin, 10 mg/ml aprotinin, 50 mg/ml phenylmethylsulfonyl fluoride (PMSF; CAUTION), 0.2 mM sodium orthovanadate, 100 mM sodium fluorine (CAUTION)).
• 2| Transfer each milliliter of cell suspension into a centrifuge tube, and spin the tubes at maximum speed at 4 °C for 15 min in a centrifuge.
• 3| Pool the supernatants (30 ml) and add 30 mg of the appropriate antibody (that will precipitate the target protein). Rock the immunoprecipitate at 4 °C for 1 h.
• 4| Add 0.9 ml of protein A?Sepharose slurry. Rock the immunoprecipitate at 4 °C for another 30 min.
• 5| Wash the protein A?Sepharose mixture in NETN buffer (20 mM Tris (pH 8), 1 mM EDTA, 900 mM NaCl, 0.5% NP-40). Repeat this wash five more times. Finally, wash the mixture once in NETN buffer containing 100 mM NaCl.
• 6| Remove the liquid portion of the mixture by aspiration. Add 800 ml of 1X SDS gel-loading buffer to the beads and boil for 4 min.
• Analysis of the immunoprecipitate
• 7| Load the sample into a large well of the discontinuous SDS-PAGE gradient gel and run the gel overnight at 10 mA constant current.
• 8| Visualize the protein bands by staining with Coomassie blue, excise the band of interest from the gel and place it in a centrifuge tube. Wash the gel slice twice for 3 min each in 1 ml of 50% acetonitrile ( CAUTION).
• 9| Digest the protein with trypsin while it is still in the gel, and electroelute the peptides.
• (i) Remove the gel slice to a clean surface, and allow it to partially dry.
• (ii)Add 5 ml of trypsin digestion buffer and 2 ml of trypsin solution.
• (iii)After the gel absorbs the trypsin solution, add 5-ml aliquots of trypsin digestion buffer until the gel slice regains its original size.
• (iv)Place the gel slice in a centrifuge tube, immerse it in trypsin digestion buffer and incubate it for 4 h at 30 °C. Stop the reaction by addition of 1.5 ml of 0.1% trifluoroacetic acid.
• 10| Fractionate the peptides by narrow-bore high-performance liquid chromatography. Subject the collected peptides to automated Edman degradation sequencing on an ABI 477A or 494A machine.
• two-hybrid assays
• yeast two-hybrid (YTH) assay. Each of the 2 putative interacting proteins is fused to one of the 2 functionally distinct domains of the transcription factor Gal4. The "bait" comprises a protein fused to the Gal4 DNA-binding domain, and the "prey" comprises a protein fused to the Gal4 transcriptional activation domain. Physical interaction between bait and prey brings the DNA-binding domain and the activation domain of Gal4 into close proximity, thereby reconstituting a transcriptionally active Gal4 hybrid. Gal4 activity can be assayed by the expression of reporter genes and selectable markers. However, its use is limited by host organism and nuclear localization requirements, and a tendency to detect indirect interactions (false positives)^ref.
• detection of protein-protein interactions that require a specific post-translational modification : modified YTH system, using the acetylation of histones and the phosphorylation of the carboxyl terminal domain (CTD) of RNA polymerase II as test modifications. In this tethered catalysis assay, constitutive modification of the protein to be screened for interactions is achieved by fusing it to its cognate modifying enzyme, with the physical linkage resulting in efficient catalysis. This catalysis maintains substrate modification even in the presence of antagonizing enzyme activities. A catalytically inactive mutant of the enzyme is fused to the substrate as a control such that the modification does not occur; this construct enables the rapid identification of modification-independent interactions. Proteins with links to chromatin functions that interact with acetylated histones, and proteins that participate in RNA polymerase II functions and in CTD phosphorylation regulation that interact preferentially with the phosphorylated CTD have been indentified^ref
• bacterial two-hybrid assay has eliminated localization requirements and simplified many technical aspects of the procedure^ref.
• protein fragment-reassembly assay, which then directly report interactions. A suitable reporter protein is dissected at the genetic level, and the fragments are fused to bait and prey, which are then coexpressed in vivo. Bait and prey interaction brings the reporter fragments together, facilitating reassembly of the active reporter protein, giving a direct readout of the association. This method has been demonstrated for dihydrofolate reductase^{ref1, ref2}, ubiquitin^ref and the green fluorescent protein (GFP)^ref from Aequorea victoria. We recently described improvements to the original screen based on the reassembly of the GFP enhanced-stability mutant sg100 in Escherichia coli. Our system, presented in the protocol that follows, consists of 2 plasmid vectors for the independent expression of fusions with N- and C-terminal fragments of GFP, and allows for simple visual detection of protein-protein interactions with a K_D as weak as 1 mM^ref.
• materials
• reagents
• plasmid vectors pET11a-link-NGFP and pMRBAD-link-CGFP and positive control vectors pET11a-Z-NGFP and pMRBAD-Z-CGFP are available upon request

GFP-fragment reassembly vectors and cloning linker sequences.

 (a) Oligonucleotide linker sequences for amplification of target bait and prey DNA. NGFP in-frame fusions are made through the XhoI site in the forward primer sequence. The reverse primer should contain a stop codon followed by a BamHI site. Cloning into pMRBAD-link-CFGP requires a forward primer in frame with the ATG in the NcoI site. Fusion to CGFP is achieved through the in-frame AatII site primer sequence. The symbol (xxx) represents codons of the template gene to be amplified and cloned into the vectors. 6 codons (18 bp) are usually sufficient to obtain specific amplification of most target sequences. The 6-base extension (5'-aataat) is required for efficient cutting by restriction enzymes. (b) Cloning linker sites and key features of the GFP-fragment vectors. pET11a-link-GFP contains the ampicillin resistance marker (ApR) and ColE1 origin of replication (ori). The XmaI site in the linker is lost during cloning; thus, digestion with XmaI can be used to eliminate the background that results from religation. Fusion expression is achieved in a BL21 DE3 host with the addition of IPTG, which results in transcription from the T7 promoter (PT7). The rop gene product regulates the copy number of plasmids containing the ColE1 origin of replication, and the lacI gene product, the Lac repressor, reduces leaky expression in DE3 bacterial hosts. pMRBAD-link-GFP contains the kanamycin resistance gene (KnR) and p15A origin of replication. The linker SphI site is lost during cloning; thus, digestion with SphI can be used to eliminate the background that results from religation. Fusion expression is under the control of the PBAD promoter (regulated by the product of the araC gene) and is induced with L-(+)-arabinose. Note that the resulting GFP fusions have different orientations relative to the GFP fragment, which may affect the likelihood of successful GFP reassembly^ref.
• primers for cloning into pET11a-link-NGFP and pMRBAD-link-CGFP (Fig. 1)
• selective media: LB containing 100 g/ml ampicillin and LB containing 35 g/ml kanamycin
• restriction enzymes: NcoI, BamHI, AatII, XhoI, XmaI and SphI (New England Biolabs (NEB))
• thermostable DNA polymerase (for example, Deep Vent Polymerase, NEB)
• dNTP solution (10 mM)
• calf intestinal alkaline phosphatase (NEB)
• T4 DNA ligase (NEB)
• E. coli strains DH10B and BL21 (DE3), prepared as competent cells (preferably electrocompetent)
• sequencing primers for pET11a-link-NGFP (T7 terminator primer 5'-tatgctagttattgctcag-3') and pMRBAD-link-CGFP (5'-ctactgtttctccatacccg-3')
• ExoSAP-IT (USB)
• screening medium: for 250 ml LB agar (for about ten plates) add 250 ml of 10 mM IPTG, 2.5 ml of 20% arabinose, 250 ml of 100 mg/ml ampicillin (100 mg/ml) and 250 ml of 35 mg/ml kanamycin (35 mg/ml). Sterilize all additives by passing through a 0.2 mm filter.
• lysis buffer: 50 mM Tris HCl, pH 8.0, 300 mM NaCl, 10 mM imidazole, 0.1% Triton X-100, 1 mg/ml lysozyme, 0.5 mM CaCl2, 5 mM MgCl₂
• nucleases: DNase and RNase (Sigma)
• Ni-NTA agarose (Qiagen)
• wash buffer: 50 mM Tris HCl, pH 8.0, 300 mM NaCl, 20 mM imidazole
• equipment
• thermal cycler programmed with the desired amplification protocol
• handheld long-wave UV lamp (365 nm)
• disposable standing column (Bio-Rad)
• procedure
• preparation of vectors encoding the fusion proteins
• 1. To prepare the cloning vectors, transform separately pET11a-link-NGFP and pMRBAD-link-CGFP into an E. coli strain that gives high-quality plasmid DNA (for example, DH10B or XL1-Blue) and plate onto selective agar media. Select for cells carrying the pET11a constructs in the presence of 100 g/ml ampicillin and for those carrying pMRBAD constructs with 35 g/ml kanamycin. Prepare plasmid DNA using standard miniprep protocols. pMRBAD-link-CGFP transformations must be incubated with shaking at 37 °C for 1 h before they are plated on selective media. pMRBAD-link-CGFP uses the kanamycin resistance gene as a selectable marker. All transformations involving pMRBAD-CGFP constructs (steps 1 and 4) must be followed by incubation, with shaking at 37 °C for at least 1 h, before plating on kanamycin-containing media. If a low plasmid yield is obtained for pMRBAD-link-CGFP, vector origin of replication is p15A (low copy number). Miniprep yield is usually sufficient for sequencing. Plasmid copy number can be increased by adding chloramphenicol (150 mg/ml) after overnight growth and continuing growth for 8 h. For larger-scale vector purification (midi- and maxiprep), DNA preparation by alkaline lysis/PEG precipitation is best.
• 2. To prepare vectors pET11a-link-NGFP and pMRBAD-link-CGFP for cloning, set up the following reactions for double digestion:
• reaction 1:
• pET11a-link-NGFP : 1 mg
• NEB buffer 4 BamHI : 5 ml
• 100x BSA : 0.5 ml
• XhoI (20 units) : 1 ml
• BamHI (20 units) : 1 ml
• water to : 50 ml
• reaction 2 :
• pMRBAD-link-CGFP : 1 mg
• NEB buffer 4 : 5 ml
• 100x BSA : 0.5 ml
• NcoI (20 units) : 2 ml
• AatI (20 units) : 1 ml
• water to : 50 ml
Incubate the reactions at 37 °C for 3 h, adding 1 ml of calf intestinal alkaline phosphatase (10 units) for the final 30 min. If you get poor cloning efficiency into pET11a-link-NGFP, some have experienced problems digesting with BamHI. Ensure that the total glycerol concentration <5% and incubate reaction for a maximum of 3 h. BamHI cannot be heat inactivated. Remove by phenol/chloroform/isoamyl alcohol extraction. We recommend using pMRBAD-link-CGFP for efficiency-sensitive cloning (for example, libraries).
• 3. Purify the vector products by electrophoresis through 0.8–1% agarose and determine the concentration of vector DNA by measuring UV absorbance at 260 nm or by running an aliquot on an agarose gel. We purify the DNA by electrophoresis through Whatmanglass fiber filter paper (GF/C) onto dialysis membrane, but commercial methods (for example, gel solubilization and extraction procedures) are acceptable. Ethanol-precipitate the DNA before ligation to concentrate the sample and remove salts that can interfere with ligation. The amount of vector obtained is usually sufficient to perform 2 260-ng (0.05-pmol) ligations.
• 4. Set up an amplification reaction to prepare each of the inserts, incorporating the appropriate linker sequences (Fig. 1a) to produce in-frame GFP-fragment fusions:

For a 50 ml reaction :
• template DNA : 0.1 ng
• primer (each) : 100 pmol
• 10x amplification buffer : 5 ml
• dNTP solution : 5 ml
• thermostable DNA polymerase : 1 ml
Depending on the composition of the amplification buffer, additional Mg²⁺ may be required for template DNA above 1 kilobase (kb). Alternatively, small inserts (<150 base pairs, bp) can be prepared synthetically, by annealing 2 oligonucleotides and extending with Klenow DNA polymerase. It is critical that the digested fragments produce in-frame fusions (see Fig. 1b). It is important to consider carefully into which vector the bait and prey should be cloned. For each bait-prey experiment, two orientations are possible: NGFP-bait with prey-CGFP and NGFP-prey with bait-CGFP. Knowledge of the bait-prey complex topology, and the location of free N and C termini, is especially useful at this stage. Avoid fusions to the GFP fragments at the site of protein-protein interaction and consider optimizing the spacer length between the fusions and GFP fragments, if necessary. In the absence of structural information, both orientations of bait and prey should be tried. If you get poor cloning efficiency into pET11a-link-NGFP, some have experienced problems digesting with BamHI. Ensure that the total glycerol concentration <5% and incubate reaction for a maximum of 3 h. BamHI cannot be heat inactivated. Remove by phenol/chloroform/isoamyl alcohol extraction. We recommend using pMRBAD-link-CGFP for efficiency-sensitive cloning (for example, libraries).
• 5. Amplify the nucleic acids according to the following program:

cycle number	denaturation	annealing	polymerization
1-30	1 min at 95°C	30 s at 55°C	1 min at 72°C
last			5 min at 72°C

Times and temperatures are optimized for a 1-kb template but may need to be adapted to suit the particular reaction conditions. For example, the duration of the polymerization step should be 1 min/kb for Deep Vent DNA polymerase. If a low plasmid yield is obtained for pMRBAD-link-CGFP, vector origin of replication is p15A (low copy number). Miniprep yield is usually sufficient for sequencing. Plasmid copy number can be increased by adding chloramphenicol (150 mg/ml) after overnight growth and continuing growth for 8 h. For larger-scale vector purification (midi- and maxiprep), DNA preparation by alkaline lysis/PEG precipitation is best.
• 6. To purify the amplification products, extract each reaction once with phenol/chloroform/isoamyl alcohol (25: 24:1, v/v/v) and then with chloroform/isoamyl alcohol (29:1, v/v) and recover the DNA by precipitation with ethanol. Analyze the size and quality of the products by electrophoresis through an agarose gel.
• 7. To prepare the inserts for cloning, digest 1 mg of insert DNA of the appropriate vector as described in step 2, omitting the calf intestinal alkaline phosphatase treatment.
• 8. Set up the reactions for ligation of inserts into vectors, by mixing 0.05 pmol of vector with 0.05 pmol of insert (260 ng pET11a-link-NGFP and 150 ng pMRBAD-link-CGFP per reaction):
• doubly digested, dephosphorylated vector DNA (step 2) : 0.05 pmol
• doubly digested insert DNA (step 7) : 0.05 pmol
• 10x ligase buffer : 1.0 ml
• DNA ligase (400 units) : 0.5 ml
• water : to 10 ml
Incubate reactions at 16 °C for at least 3 h. To optimize transformation efficiencies, empty religated vector can be eliminated by digesting the reaction with 0.5 ml of restriction enzyme corresponding to the linker site removed through cloning (XmaI for pET11a-link-NGFP, SphI for pMRBAD-link-CGFP). Incubate the reaction at 37 °C for 1 h, followed by heat inactivation at 80 °C for 20 min. This additional step is especially useful when a low background is important (for example, when cloning libraries). The target sequence must not contain the linker restriction site that is removed through cloning.
• Transformation and analysis of the clones
• 9. Transform the ligated products by mixing 1 ml of each reaction (from step 8) with 50 ml electrocompetent E. coli DH10B. Transform, preferably by electroporation (2-mm cuvette at 2.5 kV), and quench with 1 ml of 2YT. Plate the pET11a-link-NGFP transformations immediately onto LB agar containing 100 mg/ml ampicillin. Incubate the pMRBAD-link-CGFP transformations with shaking (>200 r.p.m.) at 37 °C for at least 1 h, then plate onto selective agar medium containing 35 mg/ml kanamycin.
• 10. Select several colonies, grow each in 5 ml of 2YT medium with the appropriate antibiotic and prepare plasmid DNA from 1.5 ml of culture using a standard miniprep procedure.
• 11. Verify the presence of the insert by digesting 10 ml of each miniprep with the appropriate pair of restriction enzymes and separating the fragments by electrophoresis through an agarose gel. Alternatively, the loss of the unique linker restriction sites can also be used as a diagnostic.
• 12. Confirm that the GFP fusions are the correct ones by sequencing the clones using appropriate primers (Fig. 1a). In the specific case of pMRBAD-CGFP constructs, which we prefer for libraries, we perform colony PCR with primers 5'-tagcggatcctacctgacgc-3' and 5'-ttcgggctttgttagcagcc-3'. Reactions are performed in a 96-well plate and are cleaned up for sequencing by treatment with ExoSAP-IT at 37 °C for 15 min, followed by 80 °C for 20 min.
• 13. Transform the verified plasmid constructs into a DE3 host (we use electrocompetent BL21). When a single pair of constructs is to be checked, simultaneous transformation of both plasmids is possible, using 1–10 ng of each construct miniprep DNA in 1 ml water. Incubate the doubly transformed cells with shaking at 37 °C for 1 h and then plate them on doubly selective LB agar plates (100 g/ml ampicillin and 35 g/ml kanamycin). When the transformation efficiency is critical (for example, when one plasmid contains a library), we prepare electrocompetent cells containing the constant partner.
• Screening for GFP reassembly
• 14. To perform the screen for GFP reassembly, plate 5–10 ml of a 1:10⁴ dilution of a fresh overnight culture onto screening media to obtain individual colonies. We recommend marking the agar plate into quadrants, for control and experimental combinations (Fig. 2a). This allows for side-by-side comparison of bait and prey sets (Fig. 2b). Incubate the plates either at 30 °C overnight followed by 15–25 °C for 1–2 d, or continuously at 15–25 °C for 2–3 d. After the first day, it is advisable to seal plates with Parafilm, to prevent them from drying out.

Figure 2. (a) Schematic plate layout for screening interactions with the GFP reassembly screen.

Z^N and Z^C represent control leucine zipper constructs (positive control), and BN and PC are the bait and prey NGFP and CGFP constructs, respectively. This arrangement provides a positive control, two negative controls and an experimental (exp) quadrant. (b) Examples of GFP reassembly with leucine zipper peptides. Top, empty vectors and single leucine zipper expression combinations do not give green fluorescence. Bottom, fluorescence observed with the original NZGFP and CZGFP constructs^ref (left) and improved vectors (right). (c) GFP reassembly mediated by the TPRs of HOP7. pET11a-link-NGFP constructs contained the negative control leucine zipper peptide (Z) and the C-terminal 24 residues of Hsp70 and Hsp90. pMRBAD-link-CGFP contained TPR1, TPR2A and TPR2B from HOP. Cognate interactions (70:TPR1, 90:TPR2A) are brighter than noncognate interactions (70:TPR2A, 90:TPR1), but affinities measured in vitro by surface plasmon resonance do not directly correlate to brightness. NSB, no saturating binding. Typical induction conditions are 10 mM IPTG for pET11a-NGFP and 0.2% arabinose for pMRBAD-link-CGFP. These conditions give reproducible green-fluorescent colonies with the control leucine zipper bait and prey peptide fusions. Incubation at 15–25 °C is required for the development of green fluorescence. Incubation at 37 °C alone does not produce fluorescent colonies. If you observe no green fluorescence or poor fluorescence when screening colonies for presence of the reassembled GFP, it may be due to
• poor expression of fusions. Check for expression of GFP fusions on SDS-PAGE. Vary concentrations of inducer to obtain roughly stoichiometric levels of expression. 0–20 mM IPTG and 0.02–0.2% arabinose is useful. Induction of pET11a-NGFP fusions may not be necessary, because the T7 promoter is leaky. Addition of >0.2% arabinose does not significantly improve expression of CGFP fusions.
• configurational incompatibility between bait and prey GFP fusions. GFP fragments could interfere with bait-prey interaction. Alternatively, bait and prey may interact, but the GFP fragments may be unable to reassemble. Try increasing bait-prey-GFP spacer length (for example, through multiple GlyGlySer insertions) or swapping bait-prey vector orientation. We have successfully used spacers up to 15 amino acids in length.
• bait or prey misfolded. The analyte protein most likely to be an independently structured domain should be placed on pMRBAD-link-CGFP vector, so that the unstructured CGFP fragment follows the folded domain. We typically display short peptides and unstructured motifs as NGFP fusions. Exploring alternative GFP dissection and fragment fusion strategies may be necessary.
• low reassembly efficiency. Factors such as expression level and solubility could lead to visually undetectable fluorescence in vivo. Perform expression in liquid culture and then use the NGFP His-tag to simultaneously purify and concentrate the soluble, reassembled material for quantitation in vitro. This method is very sensitive and should reveal if any reassembly (and therefore bait-prey interaction) is taking place.
• 15. Observe the plates with a handheld long-wave UV lamp (365 nm) to visualize colonies that exhibit fluorescence (thus having properly reassembled GFP). Green fluorescence under long-wave ultraviolet illumination (365 nm) typically develops after the second day and does not improve beyond 4 days. Suitable safety measures (safety glasses, gloves) should be employed when using UV light. Most brands of disposable plastic Petri dishes (for example, Falcon 1029) transmit UV light poorly. Visualization must be performed on open plates, and care should be taken in their handling. If you observe no green fluorescence or poor fluorescence when screening colonies for presence of the reassembled GFP, it may be due to :
• poor expression of fusions. Check for expression of GFP fusions on SDS-PAGE. Vary concentrations of inducer to obtain roughly stoichiometric levels of expression. 0–20 M IPTG and 0.02–0.2% arabinose is useful. Induction of pET11a-NGFP fusions may not be necessary, because the T7 promoter is leaky. Addition of >0.2% arabinose does not significantly improve expression of CGFP fusions.
• configurational incompatibility between bait and prey GFP fusions. GFP fragments could interfere with bait-prey interaction. Alternatively, bait and prey may interact, but the GFP fragments may be unable to reassemble. Try increasing bait-prey-GFP spacer length (for example, through multiple GlyGlySer insertions) or swapping bait-prey vector orientation. We have successfully used spacers up to 15 amino acids in length.
• bait or prey misfolded. The analyte protein most likely to be an independently structured domain should be placed on pMRBAD-link-CGFP vector, so that the unstructured CGFP fragment follows the folded domain. We typically display short peptides and unstructured motifs as NGFP fusions. Exploring alternative GFP dissection and fragment fusion strategies may be necessary.
• low reassembly efficiency. Factors such as expression level and solubility could lead to visually undetectable fluorescence in vivo. Perform expression in liquid culture and then use the NGFP His-tag to simultaneously purify and concentrate the soluble, reassembled material for quantitation in vitro. This method is very sensitive and should reveal if any reassembly (and therefore bait-prey interaction) is taking place.
• Purification of the reassembled GFP complex
• 16. If fluorescent cells do not appear, select a candidate colony from those identified in step 15, and prepare a 50-ml culture in LB, supplemented with ampicillin and kanamycin but without IPTG or arabinose. Prepare also positive (containing leucine zipper peptide) and negative (containing the empty vectors) control cultures for quantitative comparison. Grow the cultures at 37 °C overnight in a shaking incubator Alternatively, bacteria from an agar plate may be recovered using a plate spreader and 1 ml of TN buffer (50 mm Tris HCl, pH 8.0; 300 mm NaCl). The pET11a-link-NGFP vector incorporates an N-terminal hexahistidine tag. Because GFP reassembly is effectively irreversible, and unassembled GFP-fragment fusions are typically insoluble, the reassembled fusion can be purified and concentrated in a single step from the soluble fraction of cell lysate.
• 17. The next day, move the flasks to 15–25 °C and add IPTG and arabinose to the final induction concentrations (see step 14). Continue shaking for a further 2–4 d at 15–25 °C.
• 18. Harvest the cells by centrifugation, and resuspend them in 2 ml of lysis buffer. Add 1 l each of DNase and RNase and incubate the lysate on ice for 30 min.
• 19. Split the lysate into 1 ml aliquots and centrifuge at 4 °C for 30 min at maximum speed in a benchtop microcentrifuge.
• 20. Combine the superatant with 200 ml Ni-NTA agarose slurry and mix gently at 4 °C for 1 h. Wash the resin in a disposable standing column with 2 ml of wash buffer and then elute His-tagged protein with wash buffer plus 300 mM imidazole. Fluorescence per cell culture absorbance unit (600 nm) can be assessed quantitatively with an emission scan between 475 and 525 nm (excitation = 468 nm). Reassembled GFP variant sg100 has a emission at 505 nm.
Protein fragment-reassembly screens based on GFP reassembly offer several key advantages. First, detection of interactions is direct (green fluorescence) and does not require specific subcellular localization or a chemical substrate. Because the reassembly of GFP is effectively irreversible, fluorescence tends to accumulate over time, which probably aids in the detection of weak interactions. We have successfully used the screen to detect the values of K_D between 1 mM and 1 mM. GFP also makes the screen amenable to fluorescence-activated cell sorting (FACS) procedures, which simplifies the selection of positive interaction pairs. Second, GFP expression has been documented in a wide variety of organisms, in nearly every cell type and in most subcellular locations^{ref1, ref2}. GFP tagging often has no effect on protein function, and subcellular localization does not generally affect GFP chromophore maturation. Protein-protein interactions can therefore be studied in their native biological environment. Similar reassembly schemes have also been demonstrated for the related yellow and cyan fluorescent proteins, with alternative dissection and fusion sites within the molecule^{ref1, ref2}. Reassembly to form hybrid fluorescent proteins, with unique spectral emission properties, opens the prospect of simultaneous detection of multiple tissue-specific expression patterns within an organism^ref.
• glutathione-S-transferase (GST) fusion proteins have had a range of applications since their introduction as tools for synthesis of recombinant proteins in bacteria^ref. Typically, GST pull-down experiments are used to identify interactions between a probe protein and unknown targets and to confirm suspected interactions between a probe protein and a known protein^{ref1, ref2}. The probe protein is a GST fusion, whose coding sequence is cloned into an isopropyl-b-D-thiogalactoside (IPTG)-inducible expression vector. This fusion protein is expressed in bacteria and purified by affinity chromatography on glutathione-agarose beads. Target proteins are usually lysates of cells, either labeled with [³⁵S]methionine or unlabeled, depending on the method used to assay the interaction between the target and the probe. The cell lysate and the GST fusion protein are incubated together with glutathione-agarose beads. Complexes recovered from the beads are resolved by SDS-PAGE and analyzed by western blotting, autoradiography or staining^ref.
• precleaning the cell lysate
• incubate the cell lysate with 50 ml of a 50% slurry of glutathione-agarose beads and 25 mg of GST for 2 h at 4 °C with end-over-end mixing. The amount of lysate needed to detect an interaction is highly variable. Start with a volume of lysate equivalent to 1 x 10⁶–1 x 10⁷ cells.

Because the aim of the experiment is to compare GST with a GST fusion protein, it is necessary to prepare enough precleared lysate for each reaction. Efficient mixing of reagents is the key to success and is best achieved if the reaction is carried out in a reasonable volume: 500–1,000 ml is a good starting point. Typically a cell lysate is used in which the proteins are labeled with ³⁵S. It is, however, possible to use unlabeled cell lysates, depending on the goals of the experiment and the desired detection method.
• centrifuge the mixture at maximum speed for 2 min at 4 °C in a microcentrifuge.
• transfer the supernatant (the precleared cell lysate) to a clean microcentrifuge tube.
• probing the cell lysate
• set up 2 microcentrifuge tubes, each containing equal amounts of precleared cell lysate and 50 ml of glutathione-agarose beads. To one tube add 10 mg of GST protein; to the other tube add 10 mg of the GST fusion probe protein. The amount of probe and control protein added should be equimolar in the two reactions (that is, the final molar concentration of GST should be the same as that of the GST fusion probe protein).
• incubate the tubes for 2 h at 4 °C with end-over-end mixing, and centrifuge the samples at maximum speed for 2 min in a microcentrifuge.
• save the supernatants at 4 °C in clean microcentrifuge tubes. These samples will be analyzed by SDS-PAGE in step 9.
• wash the beads four times with 1 ml of ice-cold GST lysis buffer. Centrifuge the tubes at maximum speed for 1 min in a microcentrifuge. Discard the supernatants.

GST lysis buffer consists of 20 mM Tris-Cl (pH 8.0), 200 mM NaCl, 1 mM EDTA (pH 8.0), 0.5% Nonidet P-40, 2 mg/ml aprotinin, 1 mg/ml leupeptin, 0.7 mg/ml pepstatin and 25 mg/ml phenylmethylsulfonyl fluoride (PMSF).
• optional: elute the GST fusion protein and any proteins bound to it by adding 50 l of 20 mM reduced glutathione in 50 mM Tris-Cl (pH 8.0) to the beads. Centrifuge the tubes at maximum speed for 2 min in a microcentrifuge.
• detecting interacting proteins : analyze the beads (from step 7) or the eluted proteins (from step 8) by SDS-PAGE. Detect the proteins associated with the GST fusion protein by one of the following means.The method of detecting proteins associated with the GST fusion protein will depend on whether or not the cell lysate was radiolabeled and on the goal of the experiment.
• if the goal is to detect all of the ³⁵S-labeled proteins associated with the fusion protein, dry the gel on a gel dryer and expose it to X-ray film to produce an autoradiograph.
• if the goal is to detect specific associated proteins, transfer the proteins from the SDS-polyacrylamide gel to a membrane and perform immunoblotting.
• if the goal is to determine the sizes and abundance of proteins associated with the fusion protein from a nonradioactive lysate, stain the gel with Coomassie blue or silver nitrate.
• protein microarrays :
• MHC-peptide arrays : the identification and quantification of T cells specific for a particular antigen is a highly effective means for characterizing a subject's reaction to an immunogenic stimulus, such as from a pathogen or a vaccine. Unfortunately, this can be a daunting process; some effective techniques exist, such as cell proliferation assays, but these can be time-consuming and are ineffective for the conduct of large-scale experiments. In the body, antigens are presented to the immune system as peptides bound to cell-surface major histocompatibility complex (MHC) proteins. In 2003, Stanford University investigators developed an array-based system that used such immobilized MHC-peptide complexes to bind and isolate T cells that target a specific epitope^ref, demonstrating new promise for high-throughput T-cell analysis. Another step allows investigators not only to isolate specific T-cell populations, but also to determine the extent and the nature of activation in response to antigen binding^ref. Each array spot contains MHC complexed with different peptides, alongside costimulatory antibodies; however, the spots also include antibodies specific for cytokines released by T cells following activation. After binding cells to the array, the results of the experiment are visualized with fluorescent antibodies that also recognize those cytokines. The actual advance here is using the capture antibodies together with the MHCs, so that they can not only attract, bind and count the cells, but also look at their functional responses on the microarray. Because the detection process is specific for activation, only functional interactions should be observed; furthermore, by simultaneously screening for several different cytokines, one can very specifically characterize the nature of T-cell activation.


(a) Each spot contains immobilized MHC proteins (red), which present different peptides, and costimulatory antibodies (not shown). (b) T cells that recognize a given epitope will bind the array and release cytokines (yellow), which are bound by capture antibodies on the array. (c) Fluorescently tagged antibodies (blue) against these same cytokines allow detection.
The sensitivity appears to be enough to pick up cells at a frequency that you might expect to find them in blood from a person who's had a vaccine or been exposed to an infectious agent.. The advantage of this technology in general is really that they're parallelizing and miniaturizing existing assays, so they can be done on a larger scale. Initial experiments using a nonfluorescent, precipitatable detection substrate also suggest that even facilities without expensive array-scanning equipment, could avail themselves of this high-throughput diagnostic tool
• photo-cross-linking interacting proteins with a genetically encoded benzophenone : a major challenge in understanding the networks of interactions that control cell and organism function is the definition of protein interactions1, 2, 3, 4. Solid-phase peptide synthesis has allowed the photo-cross-linkable amino acid p-benzoyl-L-phenylalanine (pBpa) to be site-specifically incorporated into peptide chains, to facilitate the definition of peptide-ligand complexes5, 6. This method, however, is limited to the in vitro study of peptides and small proteins. An innovative development allows the incorporation of a site-specific photo-cross-linker into virtually any protein that can be expressed in Escherichia coli, thereby promoting in vivo or in vitro cross-linking of proteins7, 8, 9. The method relies on an orthogonal aminoacyl tRNA synthetase-tRNA^CUA pair that incorporates pBpa at the position encoded by the amber codon (UAG) in any gene transformed into E. coli7. The system described in this protocol uses 2 plasmids: a p15A-based plasmid to express the orthogonal tRNA and synthetase pair (pDULE) and a second plasmid containing an amber mutant of the gene of interest. To produce the photo-cross-linker?containing protein, cultures of E. coli carrying both plasmids are grown in the presence of the unnatural amino acid. To photo-cross-link the protein to its binding partner in vivo or in vitro, cells or purified proteins, respectively, are exposed to UV light


(a) The chemical structure of pBpa. C-H bonds within 3 Å of the carbonyl oxygen are targets for cross-linking. (b) pDULE-pBpa plasmid. The pBpa-specific aminoacyl-tRNA synthetase gene is located between the lpp promoter and rrnB terminator. The tRNACUA gene is located between an lpp promoter and an rrnC terminator. The plasmid contains a p15A origin and a tetracycline resistance marker
The steps to express a protein with a site-specifically incorporated pBpa in E. coli :
• materials
• reagents
• Talon metal affinity resin (BD Biosciences)
• Coomassie (Bradford) Protein Assay kit (Pierce)
• E. coli strain DH10B, prepared as electrocompetent cells
• elution buffer: 50 mM sodium phosphate, 300 mM sodium chloride, 150 mM imidazole (pH 7)
• heavy metal-containing glycerol minimal medium^ref prepared as described in the text (Step 5)
• InVision His-tag In-gel stain (Invitrogen)
• Lysis buffer: 1 mg/ml lysozyme, 50 mM sodium phosphate, 300 mM sodium chloride (pH 7)
• PAGEr Gold precast polyacrylamide gels, 4?12% Tris-glycine, 9 x 10 cm, 1-mm thick (Cambrex)
• phosphate-buffered saline (PBS): 10 mM sodium phosphate, 140 mM sodium chloride (pH 7.2)
• pBpa (Bachem) solublized in 1 M sodium hydroxide solution (Step 6)
• Petri dish (polystyrene, tight lid, 47-mm; Fisher)
• plasmid vectors: pTrcHis (Invitrogen); pDULE-pBpa (Fig. 1b),
• pDULE-Tyr (available on request from Peter Schultz (Schultz@scripps.edu)
• poly-Prep chromatography columns (0.8 x 4 cm; Bio-Rad)
• polystyrene 96-well plate (flat, low evaporation, 0.1?2 l; Costar)
• selective medium: 2X YT
• SOC medium
• wash buffer: 50 mM sodium phosphate, 300 mM sodium chloride (pH 7)
• isopropyl--D-thiogalactoside (IPTG)
• PD10 columns (Amersham)
• equipment
• light panel, camera and Digigenius (SYNGENE) software for imaging and quantifying silver-stained SDS-PAGE gels
• sonic dismembrator (model 100; Fisher)
• UV filters: Pyrex recrystallization dish (3-mm thick) or Petri dish (polystyrene, tight lid, 47-mm; Fisher)
• UV lamp: Rayonet RPR-100 chamber photoreactor, equipped with 16 350-nm bulbs (preferred) or handheld long-wavelength UV lamp such as Entela UVG1-58 at 366 nm (acceptable)
• procedure
• preparation of the mutant construct and transformation
• 1. To introduce the photo-cross-linkable amino acid pBpa into your protein of interest (your favorite protein; YFP), clone the gene encoding this protein (your favorite gene; YFG) into a standard IPTG-inducible overexpression vector (for example, pTrcHis) to create the plasmid pTrc-YFG. The proteins should be His-tagged for purification on the Talon resin. The overexpression vector does not have to be a pTrc vector. But the vector you choose must not use a tetracycline resistance selection marker and should contain an origin of replication compatible with the p15A origin of replication. Most common expression vectors meet these criteria; common examples of compatible vectors include those derived from pUC or pBR322, such as pET vectors (Novagen).
• 2. In the sequence of YFP select amino acids for replacement with pBpa. Mutate the codon for the selected amino acid in YFG to the amber codon (TAG) using a standard oligonucleotide-directed mutagenesis method to create the plasmid pTrc-YFG-TAG. We used the QuikChange method (Stratagene), but other methods are acceptable. Ideal sites for mutagenesis will be amenable to substitution: that is, mutagenesis to pBpa will not alter protein function or expression. As pBpa is an aromatic hydrophobic amino acid, is likely that tyrosine, phenylalanine or tryptophan residues will be the most amenable to replacement with the photo-cross-linker. Because it is often not clear a priori whether an amino acid will be amenable to substitution, it is worth constructing several mutants in parallel. We recommend that the interface site also be replaced with tyrosine using pDULE-TyrRS via suppression. We have seen in several cases that this serves as very good positive control for protein production and interface disruption.
• 3. Prepare 4 different protein production strains by transforming E. coli (DH10B) with the following plasmid or pair of plasmids:
• pTrc-YFG (AmpR)
• the amber mutant, pTrc-YFG-TAG (Amp^R)
• pDULE-Tyr (TetR) and pTrc-YFG-TAG (Amp^R)
• pDULE-pBpa (TetR) and pTrc-YFG-TAG (Amp^R)
• To prepare each expression strain, mix 0.2 mg of total plasmid DNA with 50 ml electrocompetent E. coli DH10B cells and transform by electroporation (in a 2-mm cuvette at 2.5 kV). Rescue the cells in 1 ml SOC medium at 37 °C for 1 h with shaking at 250 r.p.m. Select cells carrying appropriate plasmid(s) by plating onto selective LB agar medium containing 100 mg/l ampicillin and 25 mg/l tetracycline, as appropriate (if the expression vector has a marker other than the ampicillin resistance gene, substitute with the appropriate antibiotic). For delivering 2 plasmids, we find it most convenient to perform a cotransformation into DH10B electrocompetent cells and select for transformants containing both plasmids on rich-medium agar plates with both antibiotics. If colonies do not grow after cotransformation, cotransformation with both plasmids and direct selection for colonies that are both ampicillin and tetracycline resistant will have a lower transformation efficiency than transformation with single plasmids. It is therefore advisable to perform sequential transformation. First transform with pDULE plasmids and select on rich-medium agar plates containing tetracycline; then make competent cells, and transform with the pTrc-YFG-TAG plasmid into the cells containing the pDULE plasmid.
• 4. For each transformed strain, select a fresh colony grown on a rich-medium agar plate containing the appropriate antibiotic(s) and grow at 37 °C to saturation in 5 ml of 2 YT medium with 100 mg/l ampicillin and 25 mg/l tetracycline, as appropriate.
• expression of mutant protein containing pBpa
• 5. for each protein studied, 4 control expression trials should be conducted in parallel with the expression of the protein of interest as below :

expression trial	(1) native protein (positive control)	(2) background UAG (negative control)	(3) Tyr-containing protein (positive control)	(4) no pBpa control (negative control)	(5) pBpa-containing protein
YFP expression vector	pTrc-YFG	pTrc-YFG-TAG	pTrc-YFG-TAG	pTrc-YFG-TAG	pTrc-YFG-TAG
pDULE vector	none	none	pDULE-Tyr	pDULE-pBpa	pDULE-pBpa
pBpa	none	none	none	none	1 mM
antibiotics	ampicillin (50 mg/ml)	ampicillin (50 mg/ml)	ampicillin (50 mg/ml) and tetracycline (2.5 mg/ml)	ampicillin (50 mg/ml) and tetracycline (2.5 mg/ml)	ampicillin (50 mg/ml) and tetracycline (2.5 mg/ml)

For each expression trial, prepare one flask containing 200 ml of glycerol minimal medium supplemented with heavy metals as follows:
• 1.25% glycerol : 160 ml (autoclaved)
• 5X M9 salts : 40 ml (autoclaved)
• leucine (4 mg/ml) : 2 ml (autoclaved)
• CaCl₂ x 2H₂O (20 mg/ml) : 200 ml (autoclaved)
• MgSO₄ (0.12 g/ml) : 200 ml (autoclaved)
• D-biotin (5 mg/ml) : 40 ml (sterile-filtered)
• thiamine-HCl (5 mg/ml) : 40 ml (sterile-filtered)
• heavy metal stock solution : 200 ml (see recipe below)
• water : to 200 ml (autoclaved)
To prepare 1 l of 1,000X heavy-metal stock solution, combine the following metals in 1 M HCl, stir overnight at 15?25 °C, and then filter (through a 0.2-mlm filter) to remove insoluble ingredients^ref. This stock solution is stable at 15?25 °C.
• MoNa₂ x 2H₂O : 500 mg
• CoCl₂ 250 mg
• CuSO₄ x 5H₂O : 175 mg
• MnSO₄ x H₂O : 1 g
• MgSO₄ x 7H₂O : 8.75 g
• ZnSO₄ x 7H₂O : 1.25 g
• FeCl₂ x 4 H₂O : 1.25 g
• CaCl₂ x 2 H₂O : 2.5 g
• H₃BO₃ : 1 g
• 1 M HCl : to 1 l
• 6. Prewarm each flask of medium at 37 °C for 1 h with shaking at 250 r.p.m. Add the antibiotics and pBpa as described in Table 1. To obtain a final concentration of 1 mM pBpa (in flask #5), dissolve 54 mg of pBpa in 220 ml of 1 M sodium hydroxide, then add to 200 ml of prewarmed medium to give a final concentration of 1 mM. Note that the concentration of antibiotics is reduced in these cultures to maximize protein expression (50 mg/ml ampicillin and 2.5 mg/ml tetracycline). Use freshly prepared amino acid stock. The medium must be prewarmed; otherwise the amino acids may come out of solution. We occasionally observe amino acids precipitating out of solution on addition, but this has no effect on protein expression.
• 7. Centrifuge 1 ml of each of the saturated cell cultures (from Step 4) for 10 min at 1,000g and remove 0.75 ml of the supernatant. Resuspend the cells in the remaining medium (0.25 ml) and add each concentrated culture to the appropriate flask as described in Table 1. Prepare 2 1-ml aliquots of concentrated cultures of the fourth strain (transformed with both pDULE-pBpa plasmid and pTrc-YFG-TAG) for trials 4 and 5; one of these should be added to the fourth flask, the other to the fifth flask containing 1 mM pBpa (Table 1).
• 8. Incubate the cultures at 37 °C with shaking at 250 r.p.m. until they reach OD₆₀₀ = 0.6-0.8 (18-20 h for the control strains); then induce protein expression by adding IPTG to a final concentration of 1 mM (48 mg of IPTG to each 200-ml culture). Continue incubating at 37 °C for an additional 20 h with shaking at 250 r.p.m.; pellet each culture in four 50-ml aliquots by centrifugation at 5,000g. When growing multiple cultures in parallel, it is common for them to double at slightly different rates.
These cell pellets may be stored at -80 °C for use in Steps 9, 14 and 18.
• testing protein expression
• 9. Before setting up the cross-linking experiments, purify protein from each of the five cultures, using the BD Talon Affinity Resin Native Purification protocol VI B, which is summarized in Steps 9?12. Resuspend one pellet from 50 ml of cells of each culture (Step 8) in 1 ml of lysis buffer and place on ice for 20 min. Sonicate the samples at 4 °C until the viscosity is visibly reduced and then centrifuge at 20,000g at 4 °C for 30 min.
• 10. Prepare the Talon resin by suspending 150 ml of the resin (bed volume) in 1.5 ml wash buffer (ten times the bed volume). Centrifuge the resin at 700g, remove and discard supernatant, and repeat the wash process twice. Prepare 5 aliquots of resin, one for each lysate. Note that Steps 11 and 12 deal with purification of one sample.
• 11. Add the cleared lysate from Step 9 to 150 ml of washed Talon resin, and gently mix at 15-25 °C for 20 min. Centrifuge the samples at 700g to pellet the resin, and then carefully remove and discard the supernatant.
• 12. Resuspend the resin in 1 ml of wash buffer, and transfer the sample to a Poly-Prep chromatography column. Allow the column to drain by gravity flow, and wash with 5 ml of wash buffer. Elute the protein in 2.5 ml of elution buffer.
• 13. Analyze an aliquot of each sample by SDS-PAGE. The results of purification from cell lysates of strains carrying our control plasmids are shown in Figure 3. If little or no mutant protein is produced, this may have several causes. Analyze the controls to determine which of the following applies :
• the mutant protein may be toxic, so that the culture may not reach an OD₆₀₀ of 0.6. Grow cells in medium containing 2% glucose instead of glycerol, pellet when culture reaches an OD₆₀₀ of 0.6 and resuspend in glycerol-based medium containing IPTG. Take aliquots at 1-h time intervals and visualize protein expression as described (Step 9). If toxicity is still a problem, use another p15A-compatible expression vector such as the arabinose-inducible pBAD system (Invitrogen) for producing the mutant protein.
• the mutant pBpa- and tyrosine-containing mutant proteins are poorly expressed : suppression may decrease protein yield relative to the wild-type protein. The nucleotides proximal to the amber codon can have an effect on suppression efficiency^ref. If a C or T directly follows the amber codon in the gene, change this nucleotide to A or G, and repeat the expression experiment with the new mutant protein.
• the mutant pBpa-containing protein is poorly expressed, but the tyrosine-containing protein expresses well : there is a structural problem with incorporating pBpa. Try incorporating p-azido-L-phenylalanine at the same sites using pDULE-pAz. Alternatively, substitute another amino acid in the protein with pBpa.
The yield of proteins containing pBpa produced with the pDULE plasmid may vary. Expression of protein containing pBpa at several sites in the protein interface of 3 different homodimers gave yields of 6-26 mg/l pure pBpa-containing protein from the medium described (I.S.F. and R.A.M., unpublished observations). The protein yield normally reflects the production levels of native protein in this medium. Tyrosine-containing protein yields from pDULE-Tyr are often lower than both the native and pBpa-containing protein yields.
• photo-cross-linking of proteins in vivo
• 14. Resuspend a cell pellet from the third and the fifth cultures (Step 8), each in 30 ml of PBS. Centrifuge immediately at 3,000g and resuspend in 12 ml PBS. Place 3-ml aliquots into each of four disposable 47-mm polystyrene Petri dishes.
• 15. Place the zero time point sample at 4 °C for 2 h and place the three remaining samples in a Rayonet RPR-100 chamber photoreactor at 4 °C with the cooling fan on. Irradiate the samples: one sample for 30 min, the second for 60 min and the third for 120 min, removing each sample to 4 °C after irradiation for the time remaining until each has remained at 4 °C for 2 h, including the irradiation period. Many long-wavelength UV lamps emit a considerable amount of light at shorter wavelengths, which can damage protein samples. To minimize the damaging effects of short-wavelength UV light when using high-power lamps, we recommend filtering the output to remove light with wavelengths <280 nm. Filtering light can reduce cross-linking efficiencies because filters rarely allow 100% transmittance of long-wavelength light required to activate the cross-linker; therefore, if filters are used, some protein systems may require longer irradiation time. We recommend checking the percentage of transmittance of your filter over a range of wavelengths using a UV-visible spectrometer. First, take a background scan with the door open from 200 to 450 nm. Then, place your filter in the light path and, with the door open, scan the same range. The transmittance trace will inform you what fraction of the light is transmitted to your sample at different wavelengths. The polystyrene Petri dish lids recommended above filter out all light below 280 nm but allow 30% and 65% transmittance at 300 nm and 350 nm, respectively. Pyrex and window glass filters are more effective at removing damaging light if photodamage is a concern (3-mm Pyrex glass: 15% and 90% transmittance at 300 nm and 350 nm, respectively; 3-mm window glass: 0% and 90% transmittance at 300 nm and 350 nm, respectively). When irradiating many samples in parallel, the large chamber of the Rayonet RPR-100 photoreactor allows for each sample to receive the same amount of light. Handheld lamps are commonly used to cross-link benzophenones, but they do not deliver consistent spectral intensity over large surface areas. We have seen a consistent 20?30% decrease in cross-linking efficiencies when using the handheld Entela UVG1-58 UV lamp at 366 nm as compared to the Rayonet RPR-100 chamber photoreactor (I.S.F. and R.A.M., unpublished data). The sample size for irradiation can be varied considerably with consistent results as long as the surface area to sample size ratio is kept consistent. Be aware that long irradiations of small volumes can result in a decrease in sample volume. The type of filter does partly determine the irradiation time and amount of protein damage. Note also that UV radiation can cause serious damage to eyes and skin. UV-protective face shield and gloves must be worn if in contact with irradiation sources for cross-linking and In-gel stain visualization.
• 16. Remove each sample from the dish completely using 1 ml of PBS to aid in washing. Centrifuge 1 ml of each sample for analysis by SDS-PAGE. The remaining 3 ml can be centrifuged and stored at -80 °C for purification and analysis.
• 17. Resuspend the pellets from the 1-ml sample of irradiated cells in 50 ml of PBS and mix with 50 ml of 2X SDS load buffer. Heat the samples at 100°C for 10 min (mixing intermittently); remove cellular debris by centrifugation for 5 min at 16,000 r.c.f. Load 25-ml samples on a 4-12% Tris-glycine gel and run the electrophoresis at 120 V until the loading dye migrates out of the gel. Stain the gel with InVision His-tag In-gel stain by incubating the gel in the staining solution for 12 h. Visualize cross-linked and not cross-linked His-tagged protein with a standard DNA light box and camera as indicated by the manufacturer. Example results using the control constructs are shown in Figure 4a. If no in vivo cross-linking is observed by In-gel stain, protein (monomer or cross-linked protein complexes) may be at too low of a concentration to be easily visualized by In-gel stain in cell lysate. Purify the protein to concentrate it and assay cross-linking by silver-staining of SDS-PAGE gel. Denatured protein purification and gel analysis can be used if protein insolubility is suspected. If incorrect lamp or filter setup, check that your lamp and filter setup allows reproduction of the in vivo cross-linking experiment shown in Figure 4a. In vivo cross-linking can be monitored by many purification and detection methods. The method using InVision His-tag In-gel stain eliminates most variables that would prevent the detection of cross-linked protein. The incorporation of pBpa into proteins, as with any substitution with natural amino acids, may alter the protein's stability, thereby affecting the ease of protein purification and analysis.
• photo-cross-linking protein containing pBpa in vitro
• 18. Using pelleted cells from Step 8, purify protein as described in Steps 9-12. Exchange the buffer of the eluted protein (2.5 ml) to PBS, using a PD10 column. Measure the protein concentration using the Coomassie Protein Assay kit and dilute the protein samples for cross-linking. In the example shown in Figure 4b, we cross-linked at 60 g/ml protein.
• 19. For each time point, aliquot 200 l of protein sample into the wells of a polystyrene 96-well plate and cover with the tight polystyrene lid. Place the plate and filter into a Rayonet RPR-100 photoreactor at 4 °C and irradiate as described in Step 15 with a cooling fan. After each time point (0, 30, 60 and 120 min), remove each sample to 4 °C after irradiation for the time remaining until each has remained at 4 °C for 2 h. If no in vitro cross-linking is observed, increase the irradiation time with UV light. Increase the concentration of protein. Check that your lamp and filter setup allows reproduction of the in vitro cross-linking shown in Figure 4b. The stability or solubility of your protein when cross-linked may be compromised. Make sure to mix irradiated samples well before SDS-PAGE. Many long-wavelength UV lamps emit a considerable amount of light at shorter wavelengths, which can damage protein samples. To minimize the damaging effects of short-wavelength UV light when using high-power lamps, we recommend filtering the output to remove light with wavelengths <280 nm. Filtering light can reduce cross-linking efficiencies because filters rarely allow 100% transmittance of long-wavelength light required to activate the cross-linker; therefore, if filters are used, some protein systems may require longer irradiation time. We recommend checking the percentage of transmittance of your filter over a range of wavelengths using a UV-visible spectrometer. First, take a background scan with the door open from 200 to 450 nm. Then, place your filter in the light path and, with the door open, scan the same range. The transmittance trace will inform you what fraction of the light is transmitted to your sample at different wavelengths. The polystyrene Petri dish lids recommended above filter out all light below 280 nm but allow 30% and 65% transmittance at 300 nm and 350 nm, respectively. Pyrex and window glass filters are more effective at removing damaging light if photodamage is a concern (3-mm Pyrex glass: 15% and 90% transmittance at 300 nm and 350 nm, respectively; 3-mm window glass: 0% and 90% transmittance at 300 nm and 350 nm, respectively).

When irradiating many samples in parallel, the large chamber of the Rayonet RPR-100 photoreactor allows for each sample to receive the same amount of light. Handheld lamps are commonly used to cross-link benzophenones, but they do not deliver consistent spectral intensity over large surface areas. We have seen a consistent 20-30% decrease in cross-linking efficiencies when using the handheld Entela UVG1-58 UV lamp at 366 nm as compared to the Rayonet RPR-100 chamber photoreactor (I.S.F. and R.A.M., unpublished data). The sample size for irradiation can be varied considerably with consistent results as long as the surface area to sample size ratio is kept consistent. Be aware that long irradiations of small volumes can result in a decrease in sample volume. The type of filter does partly determine the irradiation time and amount of protein damage. Note also that UV radiation can cause serious damage to eyes and skin. UV-protective face shield and gloves must be worn if in contact with irradiation sources for cross-linking and In-gel stain visualization.
• 20. Analyze the extent of cross-linking by SDS-PAGE, which resolves the slowly migrating covalently cross-linked products from the starting proteins. Prepare samples for SDS-PAGE by combining 20 ml of each with 20 ml of 2 SDS load dye and heat at 100 °C for 10 min. Cool the samples and load 25 l directly onto a SDS-PAGE gel. Run the electrophoresis at 120 V until the loading dye migrates out of the gel, and analyze by silver staining the gel. Example results using the control constructs are shown in Figure 4b. If several low-mobility bands detected by SDS-PAGE, if you are cross-linking a homodimer, the singly cross-linked dimer is likely to migrate differently than the doubly cross-linked protein. Multiple bands may also reflect the interaction of your protein in several orientations or conformations.
• 21. To determine the percentage of cross-linking, integrate the band density corresponding to the cross-linked and not cross-linked material (we used Digigenius). Then calculate the fraction cross-linked using the equation : %cross-linking efficiency = (number of counts cross-linked) / (number of counts cross-linked + number of counts not cross-linked) x 100
The pDULE-pBpa plasmid allows the in vivo, facile, site-specific incorporation of the photo-cross-linking amino acid, pBpa, into proteins of almost any length. The amino acid is incorporated with high efficiency, fidelity and specificity. Because the pDULE-pBpa plasmid is compatible with common high-copy protein production plasmids, excellent protein yields can be obtained. With exposure of cells to light, protein-protein interactions can be cross-linked in vivo and rapidly visualized without the need for purification. Moreover, complementary studies can easily be performed in vitro (with crude cell extract or purified protein components). Site-specific cross-linking can help define a protein's binding partners and the topology of complexes. Expression of proteins containing pBpa at physiologically relevant concentrations in E. coli or eukaryotic cells^{ref1, ref2} may allow the definition of weak or transient protein interactions that can be irreversibly trapped by photo-cross-linking. Such interactions are probably not captured by the noncovalent affinity chromatography complex purification methods now used to map the protein interaction networks of cells^{ref1, ref2}.
• isolation of protein complexes
• antibody-mediated affinity purification of a tagged protein (bait) under conditions that allow interacting proteins to stay bound to one another. Usually, the complex is eluted by cleavage of the affinity tag. The components of the purified complex are then separated by size, isolated and identified by mass spectrometry.
• relative strengths, weaknesses, sensitivity, difficulty, speed, etc.. of the methods for deciphering the kinome

	technique	advantages	disadvantages
kinase => substrate	phosphorylation screening  - in vitro protein phosphorylation  - phage-based assays  - peptide or protein array screens	fast, easy and cheap  detects direct phosphorylation	artifact-prone  requires further validation
	genetic screens	many pioneering successes  not limited by sensitivity  aided by the availability of genome sequences	time consuming  not possible in all cases  does not show direct phosphorylation
	functional knockout  - specific inhibitors  - RNAi  - genetic knockout	rapid means of testing involvement of a kinase  in vivo techique	does not detect direct phosphorylation  nonspecific effects, cross-talk and feedback
	analogue-sensitive kinase	shows direct phosphorylation	specificity  not yet functional in cells
	phosphospecific substrate antibody, which detect protein phosphorylation in a sequence specific fashion, can detect specific and more general protein phosphorylation and form the basis of numerous commonly used techniques. Antibodies can be raised against phosphorylated peptides representing a specific site in a protein, or against a degenerate mix of phosphopeptides (for example, RxRxxpS/T for AKT), thus generating a broad range phosphospecific 'substrate' antiserumref. Purified antibodies raised against specific phosphorylation sites will typically recognize that particular phosphorylated site with greater affinity and specificity than an antiserum designed against a degenerate peptide mixture corresponding to the consensus phosphorylation site. Monoclonal or polyclonal antibodies against the phosphotyrosine moiety are exceptionally good; they are able to recognize phosphotyrosine with high affinity and specificity and provide an exceptionally convenient way to assay tyrosine phosphorylation in virtually any protein. Unfortunately, no equivalent high-affinity antibodies exist for phosphoserine and phosphothreonine, mainly because of the lower immunogenicity of the phophoserine and phosphothreonine side chains. The existing anti-phosphoserine and anti-phosphothreonine antibodies tend to recognize other phosphate monoesters as well and must be used with caution. These limitations are compounded in some approaches by the greater cellular abundance of phosphoserine and phosphothreonine than phosphotyrosine. An alternative to phosphospecific antibodies are phosphospecific binding domains such as SH2, PTB, BRCT, Polo box, 14-3-3, WW and FHA domainsref. These bind with variable affinity (50 nM to 100 M) to specific sequences in a phosphorylation-dependent manner. They have the potential advantage of being engineerableref and are easy to clone and thus can be conveniently used in a variety of molecular techniques. An AKT substrate has been isolated by means of a 14-3-3 proteinref and a c-Abl substrate by means of an SH2 domainref.	detects substrate phosphorylated in vivo	limited by preference of the kinase and specificity of antibody  proteins need identification by other techniques
kinase <=> substrate	physical association  - yeast two-hybrid screen - phage screen  - proteomic techniques	requires no knowledge of sequence preference of the kinase  high-throughput potential	needs other techniques to detect phosphorylation
	bioinformatics	extremely quick method for making predictions	relatively low reliability necessitates the use of other techniques  requires distinct kinase preferences
substrate => kinase	cross-linking	new, potentially poweful technique	unproven specificity  requires phosphorylation site to be known  in vitro technique
	in-gel kinase assays	convenient assay	poor renaturation of many kinases  in vitro technique

Flow chart of strategies available for the identification of either substrate or kinase.

Techniques labeled with * lead to kinase or substrate candidates that need to be further investigated and validated using alternative experimental strategies
Mass spectrometry : the ability to identify proteins and the post-translational modifications of a protein by MS is a considerable improvement on traditional biochemical techniques and presents a powerful array of tools to investigate protein phosphorylation.
• identifying phosphoproteins by MS : MS can be used with tremendous sensitivity to identify a protein unambiguously. This is useful in a variety of techniques that produce a band on an autoradiograph or a purified kinase and are often limited by the ability to identify the protein. Proteins can be identified by 'fingerprinting' where a protein is digested with trypsin and the characteristic masses of the peptides produced are accurately determined and used to search a database of predicted trypitic-peptide masses. Protein identification is also possible by tandem mass spectrometry (MS/MS)^ref. The first MS separation is used to select a particular mass species, which is then bombarded, causing fragmentation; the pattern of overlapping fragments produced allows the peptide sequence to be deduced, and this is used to search a database.
• identification of phosphorylation sites by MS : MS can be used to identify the individual phosphorylation sites on a protein^ref. A modification of the peptide fingerprinting technique can be used to identify phosphorylation of a protein. Covalent attachment of a phosphate group to a protein leads to an increase in mass of 80 Da, so the altered charge-to-mass ratio will be detectable in the peptide spectra. It may be possible to attribute the phosphorylation to a particular residue of the peptide; if not, MS/MS can pinpoint the site. Achieving peptide fragmentation in MS/MS by electron capture dissociation may lead to improved phosphopeptide sequencing^ref. When studying the phosphorylation of a protein with MS, it must be remembered that only the phosphorylation of analyzed peptides will be detected, and this should be considered before claiming to have described 'all' the phosphorylation sites of a particular protein.
• Large-scale identification of phosphorylation sites : a key to understanding the role of protein phosphorylation is the identification of in vivo phosphorylation sites. The high-throughput capabilities of MS are ideally suited to this task. Phosphorylated proteins may represent a small fraction of the total protein so enrichment is required by techniques such as IMAC44, anti-phosphotyrosine antibody enrichment^ref, chemical replacement of the phosphate group with an affinity tag such as biotin^ref or strong cationic resin exchange chromatography^ref. The phosphorylated subset of proteins can be isolated, and then these proteins can be individually identified by MS. Alternatively, the protein mixture can be digested with trypsin and the phosphopeptides enriched and identified, resulting in the identification of large numbers of phosphorylation sites from a single sample^ref. The capacity of such techniques will surely improve, ideally to the point of describing the phosphoproteome (all the phosphorylated proteins in the cell, estimated to be at least one-third of all intracellular proteins, perhaps 20,000 phosphorylated proteins in a cell with serine, threonine or tyrosine phosphorylated in the ratio 1,800:200:1). This would be an invaluable advance in understanding phosphorylation-based signaling networks. It would be equivalent to in vivo mapping of every protein in the cell and would allow many standard techniques to be used more powerfully. Phosphosite is a new database that aims to authoritatively catalog all in vivo phosphorylation.
• comparative analysis of samples by MS is possible by isotopic encoding. In stable isotope-labeled amino acids in cell culture (SILAC), cells are grown for several generations in the presence of an essential amino acid labeled with a nonradioactive isotope, so that all proteins become uniformly labeled^ref. Analysis by MS can then identify both the protein and the sample from which it originated, presenting numerous possibilities. The use of isotopes of 2 elements simultaneously allows more samples to be encoded and more detailed analyses, such as a time courses^ref. Samples can also be encoded by affinity tags^ref.
Companies offering kinome analysis products : Amersham Biosciences, AnaSpec, Applied Biosystems, BD Biosciences (Pharmingen), Biaffin GmbH,
Biocan, Biorad, BioSource International, Biotrend Chemikalien GmbH, BioVisioN AG, Calbiochem (EMD Biosciences), Cambrex Corp., CHEMICON,
Cell Signaling Technology, Cellular Genomics Inc., Clontech (EMD Biosciences), DiscoveRx, EMD Biosciences, Exalpha Biologicals Inc., GE Healthcare, Genomatix Corp., Globozymes, Imgenex, Invitrogen, Jerini AG, Kinasource, MBL International Corporation, Molecular Devices, Molecular Probes (Invitrogen Detection Technologies), New England Biolabs, Pepscan Systems, PerkinElmer, Pierce Biotechnology Inc., Phosphosolutions, Promega Corp.,
Rockland Immunochemicals for Research, Stratagene, Stressgen Bioreagents, Upstate, USBiological, Qiagen
A comparison of methods for studying gap junction communication :

properties/methods	microinjection of tracers	fluorescence recovery after photobleaching (FRAP)	dual whole cell patch clamping	local activation of a molecular fluorescent probe (LAMP)
Invasiveness	Invasive	Intense illumination and photo-damage	Invasive	Non-invasive
Cell integrity	Compromised	Maintained (if dyes are cell-permeable)	Compromised	Maintained
Temporal resolution	Low	Good	Very high	Good
Simplicity of set-up and execution	Special technique, limited number of cells	Require high-power laser	Technically demanding & time consuming	Easy, applicable to cell populations or tissues
Quantification of molecular transfer rates	Reliable only if cells recover very rapidly	Reliable if no photo-damage	Conductance does not always reflect molecular transfer, problematic in well-coupled cells from the interference of series resistance	Reliable and allows multiple measurements in coupled cell pairs
Multi-color imaging	Feasible	Difficult	Feasible	Feasible

• peptide conjugation

Care must be taken when choosing the mode of conjugation to prevent modification of the antigenic sequence. For optimum immune response, it is preferable to conjugate via a terminal residue/functional group of the peptide.
• peptide-carrier protein conjugates may occur via the following groups on the carrier protein :
• primary amines (-NH₂) groups :
• m-maleimidobenzoyl-N-hydroxysuccinimide (MBS) is a heterobifunctional cross-linking reagent that here adds to N-terminal amines to form amides. Do not use if your peptide contains Cys or Lys in positions other than the terminal ones.


• carbodiimides : they couple carboxylic acids to amines to form amides, so can be used with N-terminal amines. Do not use if your peptide sequence contains Asp, Glu or Lys in positions other than the terminal ones.
• 1-ethyl-3-(3-dimethylaminopropyl)carbodiimide hydrochloride (EDC)
• glutaraldehyde is a bifunctional reagent that adds to amines to form Schiff base polymers, so can be used with N-terminal amines. Do not use if your peptide sequence contains Cys or Lys in positions other than the terminal ones. The carrier activation reaction is stopped by adding NaBH₄.
• N- or C-terminal sulfhydryl (-SH) groups (do not use if your peptide contains Cys or Lys in positions other than the terminal ones)
• Cys =>
• sulfosuccinimidyl-4-(N-maleimidomethyl) cyclohexane-1-carboxylate (sulfo-SMCC). If there are no Cys in the sequence, it is often advantageous to add a Cys at the N- or C-terminus.
• m-maleimidobenzoyl-N-hydroxysuccinimide ester (MBS) is a heterobifunctional cross-linking reagent that here adds to thiols forms thioesters (maleimido-activated carrier).
• Tyr => bis-diazotized benzidine
• glutaraldehyde is a bifunctional reagent that adds to sulfhydryl to form Schiff base polymers. Do not use if your peptide sequence contains Cys or Lys in positions other than the terminal ones. The carrier activation reaction is stopped by adding NaBH₄.
• C-terminal carboxyl (-COOH) groups (Asp and Glu) or 5'-phosphate groups:
• carbodiimides : they couple carboxylic acids to amines to form amides. Carbodiimides work best when the N-terminus of the peptide is blocked, the peptide contains no Lys or Arg and has a free C-terminal carboxyl. The unwanted amino groups can be reversibly blocked with citraconic anhydride to prevent cross linking. Do not use this method if your peptide sequence contains Asp, Glu or Lys in positions other than the terminal ones.
• 1-ethyl-3-(3-dimethylaminopropyl)carbodiimide hydrochloride (EDC)


• ethylene diamine-activated carriers : carboxylate groups are modified with ethylene diamine inducing a rather "cationized " molecule, since the negative charged carboxylates are masked by positively charged amines resulting in a significant pI increase. So they can incorporate activated carboxylic acid functions or aldehydes with increased efficiency in comparison to native proteins.
• N- or C-terminal Trp
• glutaraldehyde is a bifunctional reagent that adds to indoles to form Schiff base polymers. Do not use if your peptide sequence contains Cys or Lys in positions other than the terminal ones. The carrier activation reaction is stopped by adding NaBH₄.
• N-or C-terminal hydroxyl-containing amino acids
• glutaraldehyde is a bifunctional reagent that adds to hydroxyls to form Schiff base polymers. Do not use if your peptide sequence contains Cys or Lys in positions other than the terminal ones. The carrier activation reaction is stopped by adding NaBH₄.
Conjugation to ligands that don't contain a classic functional group (e.g., -NH₂, -SH or -COOH) for chemical coupling is practiced coupling "active hydrogens" to a carrier protein via the Mannich reaction :

The ligand must be able to withstand heating (56°C) for 24-48 hours.
Either the peptide or the carrier can be activated first (pre-activated carriers are available commercially) and subsequently added to the carrier or the peptide, respectively. During conjugation the peptide-to-carrier ratio has to be ~ 40. The unconjugated peptide and the conjugation byproduct(s) are finally removed by dialysis cassette(s) for gel filtration. The peptide-to-protein ratio in the conjugate is determined by choosing an amino acid that is not present in the native peptide but is present in the native carrier protein and another amino acid that is present in both the native peptide and in the native carrier protein. The difference between the ratios of these 2 amino acids for the conjugate and for the native carrier shows how many copies of peptide have been conjugated per carrier protein.
• incorporation of synthetic amino acids into proteins is desirable in many instances, for example, having metal-binding or photoreactive properties or enhancing proteins for the design of novel therapeutics. A functional orthogonal pair of a tRNA and aminoacyl-tRNA synthetase that incorporated an unnatural amino acid, while not cross-reacting with any endogenous amino acids, has been designed using the Escherichia coli tyrosyl-tRNA synthetase (TyrRS) and amber suppressor tRNA_CUA pair (nonsense suppression methodology) to evolve into accepting unnatural amino acids and generated a large library of TyrRS mutants. A subsequent library of yeast cells carrying these mutants was then subjected to genetic selection involving 2 steps—a positive selection where only cells in which the TyrRS loaded the tRNACUA with any amino acid survived, followed by a negative selection in which any cells utilizing a natural amino acid, over an unnatural one, died. The 2 selections resulted yeast strains with a tRNA/synthetase pair that only utilized unnatural amino acids. Via this approach, Chin et al. independently incorporated 5 novel amino acids into the genetic code of S. cerevisiae, for example, creating a strain that incorporated an amino acid with a "benzophenone" side chain, useful as a photocrosslinker, into any mRNA that contained the amber suppressor codon.
• nonribosomal peptide synthesis (NRPS)

Nonribosomal peptides (NRP) are created on massive, assembly line-like synthetases. These enzymes are modular, comprising a series of functional units that can bind a naked amino acid, activate it as a thioester, and couple it to the growing peptide chain. Some synthetases are multimeric complexes, while others are single, massive proteins. Each module is about 1,000-1,200 amino acids long, making the enzymes enormous: a single 15,281-residue (1.7 MDa) protein, for instance, synthesizes cyclosporin, an 11-residue long immunosuppressant.
• linear NRPS, the sequence of modules acts as the template that defines the resulting peptide's sequence.
• iterative and nonlinear NRPS configurations exist, too, which generate more complicated structures.
Advantages :
• many of the NRP's modifications are performed during synthesis
• by assembly line components : where ribosomal translation is limited to the standard complement of 20 L-amino acids, nonribosomal peptides may contain unusual building blocks, including D-amino acids, methylated variants of the standard amino acids, and nonproteinogenic, hydroxylated, and glycosylated residues; more than 300 precursors are currently known. Sometimes the peptide products are hetero-cyclized, and the peptide backbone may be branched.
• by external proteins
Other modifications are added postsynthetically.
• NRPS produces several pharmacologically important compounds, including cyclosporin and the antibiotics penicillin and vancomycin.
Disadvantages : generates 2 - 48 residues-long oligopeptides
Evidence against NRPS : a genomic sequence capable of producing the peptide's sequence
Evidence for NRPS : the presence of signature modular peptide synthetase gene segments.
Web resources :
• Chemical Reviews vol. 97, no. 7, 1997
• H.D. Mootz et al., Ways of assembling complex natural products on modular nonribosomal peptide synthetases Chembiochem, 3:490-504, 2002.
• analyzing mAb specificity
• whole proteome microarrays : in 2003, several companies have compressed the entire protein-coding portion of the human genome onto a single chip--that's some 30,000 to 40,000 unique gene sequences per slide. One company has reduced its arrays to the size of microtiter-plate wells, for use in high-throughput biochip analysis. Whole proteome microarrays containing every protein for the relevant organism represents the ideal format for an assay to test antibody specificity, because it allows the simultaneous screening of thousands of proteins for possible cross-reactivity
• protein engineering :
• increasing serum half-lives of proteins
• PEGylation
• fusion to serum albumin
• ribose-binding protein (RBP), a protein with no catalytic abilities, has been transformed by computational design into triose phosphate isomerase (TIM), a highly active enzyme^ref : algorithms to predict mutations that altered RBP's layer of residues so it could bind the aldose glyceraldehyde-3-phosphate (GAP). and the ketose dihydroxyacetone phosphate (DHAP) . The authors then introduced catalytically active residues into this receptor design. First, they defined the most favorable geometrical orientations of key interactions contributing to catalysis—specifically, those of the three catalytically essential residues glutamate, histidine, and lysine with respect to the enediol intermediate. Next, the researchers used a combinatorial search algorithm to find positions for these residues that satisfy these geometrical constraints. Finally, they used their receptor design algorithm to optimize the potential active site. 14 RBP variants were designed, produced, and assayed for TIM activity. Just like TIM, the RBP variants had a mechanism to close the active site after the substrate was bound, except they used a hinge-bending mechanism. The most active variant was much less thermostable than native RBP and was stabilized with computational design by adding mutations to the protein matrix, correcting interactions with the binding surfaces it surrounded. The resulting RBP variant NovoTim1.2, which had additional surface amino acid substitutions remote from the active site, catalyzed the TIM reaction at a rate 100,000- to 1,000,000-fold more efficiently than uncatalyzed reactions. It also proved biologically active, supporting the growth of Escherichia coli under gluconeogenic conditions just as regular TIM would. To further improve the catalytic activity, the authors turned to directed evolution. NovoTim1.2 was further mutagenized randomly by an error-prone polymerase chain reaction. Because TIM is absolutely necessary for gluconeogenic growth on glycerol, the researchers plated about 100,000 cells on glycerol to select for improved mutants, and 4 colonies survived. This approach has the potential for tremendous generality. There are many reactions one would like to catalyze that have no enzyme in nature, or the enzymes that are in nature are highly fragile. So far, the approaches that have been taken are random screening or directed evolution. What they're showing here is computation, when combined with these experimental methods, now may allow one to have new starting points for evolution of highly effective and highly novel catalysts. Although the variant's efficiency parameters were about 3 orders of magnitude below that of wildtype TIM, this reduced efficiency does not diminish the achievement, considering that native TIM is a kinetically perfect enzyme with a turnover that is limited only by the diffusion-determined rate at which substrate and enzyme encounter each other. Researchers first tried to design enzymes de novo in the 1980s and came up with the first catalytic antibodies, known as abzymes. Recently, scientists at Caltech used computational design to transform the catalytically inert protein thioredoxin into an esterase^ref. However, both abzymes and the designed thioredoxin are much less active than the RBP variants transformed into TIM

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